Published online Jul 26, 2026. doi: 10.4252/wjsc.119895
Revised: April 11, 2026
Accepted: June 4, 2026
Published online: July 26, 2026
Processing time: 165 Days and 4.2 Hours
Platelets promote fracture repair by transferring mitochondria to recipient cells. Adipose-derived stem cells (ASCs) have garnered significant attention in bone regeneration due to their osteogenic differentiation potential.
To investigate the effect of platelet-derived mitochondria on ASC osteogenesis and elucidate the underlying molecular mechanism.
ASCs were isolated from Sprague-Dawley rats and characterized. ASCs were treated with platelets isolated from rat whole blood or platelet-derived mitochondria, and then evaluated for glycolysis and osteogenic differentiation. Histone lactylation and its impact on gene transcription were analyzed. A rat closed femoral fracture model was established to evaluate mitochondrial roles in vivo.
Platelets enhanced glycolysis and osteogenic differentiation of ASCs. Platelet-derived mitochondria exhibit a similar pro-osteogenic and metabolic effect, and these mitochondria were found to be internalized by ASCs. Mitochondria induced H3K18 lactylation, which enriched the promoters of AXIN2, BMPR1B, COL1A1, and OSTN, thereby promoting their transcription. 2-deoxy-D-glucose treatment or H3K18R transfection reversed the mitochondrial-induced enhancement of glycolysis and osteogenesis. Additionally, platelet-derived mitochondria amplified ASC-mediated fracture repair, an effect counteracted by 2-deoxy-D-glucose.
Platelet-derived mitochondrial transfer promotes osteogenic gene transcription via H3K18 lactylation, facilitating osteogenic differentiation and accelerating fracture healing. This study reveals a novel metabolic-epigenetic me
Core Tip: Platelets promote glycolysis and osteogenic differentiation of adipose-derived stem cells (ASCs) by transferring mitochondria. Mechanically, mitochondria facilitate H3K18 lactylation, thereby inducing transcription of AXIN2, BMPR1B, COL1A1, and OSTN. Moreover, platelet-derived mitochondria enhance the effect of ASCs on fracture healing through gly
- Citation: Pan Z, Zhang R, Du GM, Yi JH, Bao J, Liu CD. Platelet-derived mitochondria transfer accelerates fracture healing by promoting osteogenic differentiation and metabolic reprogramming of adipose-derived stem cells. World J Stem Cells 2026; 18(7): 119895
- URL: https://www.wjgnet.com/1948-0210/full/v18/i7/119895.htm
- DOI: https://dx.doi.org/10.4252/wjsc.119895
Fracture represents a common injury encountered in emergency departments. Closed reduction and internal fixation represent the cornerstone interventions for fracture management, with advancements in surgical techniques and biomaterials expediting recovery timelines[1]. However, the healing cascade is a complex, temporally orchestrated process encompassing hematoma-mediated inflammation, angiogenesis, chondrogenesis, matrix mineralization, and chronic bone remodeling[2]. Perturbations at any stage can culminate in delayed union or nonunion[3], affecting 5%-10% of patients[4], profoundly impairing quality of life and depleting healthcare resources. Therefore, exploring innovative strategies to potentiate fracture healing remains imperative.
Adipose-derived stem cells (ASCs), a mesenchymal stem cell (MSC) subset, emerge as a viable therapeutic modality for bone regeneration due to their accessibility, multilineage potential, and robust paracrine activity[5]. Notably, ASCs exhibit osteogenic plasticity, differentiating into osteoblasts to promote bone formation, positioning them as a promising candidate for augmenting fracture repair[6,7]. Nevertheless, the molecular mechanisms governing ASC osteodifferentiation remain incompletely understood.
Platelets, unique anuclear blood cells, play pivotal roles in hemostasis, inflammation, cancer metastasis, wound healing, and immune defense[8]. Activated platelets secrete growth factors, including platelet-derived growth factor, vascular endothelial growth factor, and transforming growth factor β, which orchestrate osteogenic differentiation of MSCs, endothelial cell proliferation, and fibroblast activation[2]. Leveraging these properties, platelet-rich plasma (PRP) has emerged as a widely employed therapeutic strategy to potentiate fracture healing[9]. Nevertheless, the regulatory capacity of platelets over ASC-specific osteogenesis remains poorly defined.
Mitochondria, cellular powerhouses governing metabolism and ATP production, mediate intercellular communication in physiological and pathophysiological processes[10]. Platelets, rich in mitochondria, release mitochondrial-laden extracellular vesicles and free mitochondria upon activation, contributing to systemic mitochondrial dissemination[11,12]. Accumulating evidence implicates platelet-derived mitochondria in wound healing, facilitating anti-inflammation, angiogenesis, and MSC proliferation through mitochondrial transfer[13,14]. However, the role of platelet-derived mi
The metabolic microenvironment is a critical determinant of fracture healing. During osteogenic differentiation, MSCs undergo metabolic reprogramming characterized by a shift toward glycolytic metabolism, wherein glucose is preferentially converted to lactate[15]. This metabolic adaptation likely supports the high energy demands of the healing process, requiring tight coordination between mitochondrial respiration and glycolytic flux[16]. While mitochondrial transfer-mediated metabolic reprogramming of MSCs has been implicated in wound healing[13,14], the extent to which similar mechanisms regulate metabolic shifts during fracture repair remains elusive.
This study investigates the impact of platelet-derived mitochondria on ASC osteogenic differentiation and fracture healing. Furthermore, we elucidate the regulatory role of mitochondrial transfer in ASC glycolysis and the underlying molecular mechanisms. We speculated that platelet-derived mitochondria promote glycolysis and induce H3K18 la
Sprague-Dawley (SD) rats (8-week-old, male, fifty) were purchased from CAVENS (Changzhou, Jiangsu Province, China). All rats were housed in a specific pathogen-free environment at 23-25 °C with a 12-hour light/dark cycle, providing ad libitum access to food and water. All animal procedures were conducted in accordance with the Guide for the Care and Use of Laboratory Animals. The animal protocol was approved by the Ethics Committee of Nanjing First Hospital (Approval No. DWSY-25080712).
After euthanizing the rats with isoflurane, the inguinal fat pads were collected, washed with normal saline, chopped, and digested with 0.1% type I collagenase at 37 °C for 40 minutes. Following filtration through a 75-μm cell sieve, the filtrate was centrifuged at 1600 × g for 5 minutes. The cells were washed with phosphate buffered saline (PBS) and then cultured in α-minimum essential medium (Gibco, Grand Island, NY, United States) supplemented with 10% fetal bovine serum (Gibco, Grand Island, NY, United States) and 1% penicillin/streptomycin (Gibco, Grand Island, NY, United States) under 37 °C and 5% CO2 conditions. When the cells reached 80% confluence, they were passaged. Passage 3 ASCs were used for subsequent experiments.
To evaluate the multilineage differentiation potential of ASCs, cells were cultured in rat adipogenic differentiation medium (Procell, Wuhan, Hubei Province, China). After 21 days, adipogenic differentiation was assessed via Oil Red O staining (Solarbio, Beijing, China). For osteogenic differentiation, ASCs were cultured in rat osteogenic differentiation medium (Procell, Wuhan, Hubei Province, China). After 14 days of induction, alkaline phosphatase (ALP) activity was measured using an ALP staining kit (Solarbio, Beijing, China). Osteogenic differentiation was further evaluated by Alizarin Red S (ARS) staining (Solarbio, Beijing, China) following 21 days of induction.
Stem cell surface markers were analyzed by flow cytometry. ASCs (5 × 106 cells) were washed with PBS and incubated with anti-rat PE-conjugated CD31 (Invitrogen, Carlsbad, CA, United States), anti-rat PE-conjugated CD44 (Invitrogen, Carlsbad, CA, United States), or anti-rat FITC-conjugated CD105 (Invitrogen, Carlsbad, CA, United States) at 4 °C for 30 minutes. Samples were analyzed using a flow cytometer and processed with FlowJo software.
ASCs were fixed with fixative at 4 °C for 20 minutes, rinsed with 60% isopropanol for 30 seconds, and incubated with Oil Red O staining solution for 20 minutes. After a second rinse with 60% isopropanol, the nuclei were counterstained with Mayer’s hematoxylin solution for 1 minute. Cells were then observed under a light microscope.
Following osteogenic induction and cell stimulation, ASCs were fixed with fixative at 4 °C for 30 seconds, incubated with ALP staining buffer at 37 °C for 30 minutes, and washed with distilled water. Results were observed under a light microscope for ASC identification or imaged using a digital camera for functional experiments.
Following osteogenic induction and cell stimulation, ASCs were fixed with 4% paraformaldehyde for 15 minutes. After washing with distilled water, the cells were stained with 0.2% ARS solution (pH = 8.3) at room temperature for 30 minutes. Results were observed under a light microscope (for ASC identification) or imaged using a digital camera (for functional experiments).
Blood was collected from the hearts of SD rats and immediately transferred into sodium citrate-containing tubes. After centrifugation at 200 × g for 10 minutes, the supernatant (PRP) was collected. The PRP was further centrifuged at 400 × g for 15 minutes, and the platelet pellet was collected. Platelet counts were determined using a hemocytometer.
Mitochondria were isolated from platelets using a platelet mitochondrial extraction kit (Bestbio, Shanghai, China). The isolated platelets were washed with washing buffer A, then mixed with 500 μL of pre-chilled mitochondrial extraction solution B and shaken at 4 °C for 30 minutes. After centrifugation at 2000 × g for 5 minutes, the pellet (mitochondrial-free platelets) was discarded, and the supernatant was subjected to further centrifugation at 12000 × g for 20 minutes. The resulting pellet was resuspended in 500 μL of pre-chilled mitochondrial extraction solution C and centrifuged again at 12000 × g for 20 minutes. The final pellet was resuspended in mitochondrial preservation solution D for subsequent experiments.
ASCs were seeded in 6-well plates at a density of 1 × 105 cells/well. Cells were co-cultured with 2 × 107 platelets or mi
Wild-type (WT) H3K18 and H3K18 mutant (H3K18R) were transfected into ASCs to assess the effect of H3K18 lactylation on osteogenic differentiation using Lipofectamine 2000 (Invitrogen, Carlsbad, CA, United States) following the manufacturer’s instructions. Cells were harvested 48 hours later.
Cell proliferation was assessed using the carboxyfluorescein diacetate succinimidyl ester fluorescence probe (Yeasen, Shanghai, China). ASCs (2 × 105 cells) were incubated with 500 μL carboxyfluorescein diacetate succinimidyl ester working solution at 37 °C for 30 minutes. Subsequently, cells were washed with Hanks’ balanced salt solution containing 20 mmol/L Hepes (HHBS; AAT Bioquest, Pleasanton, CA, United States) and resuspended in 500 μL HHBS. Flow cytometry was performed for analysis.
The glucose uptake assay kit (Abcam, Cambridge, MA, United States) was used to assess glucose consumption. ASCs (2 × 103 cells) were seeded in 96-well plates and serum-starved overnight. Cells were incubated with 100 μL KRPH buffer containing 2% BSA for 40 minutes. Following stimulation with 1 μM insulin for 20 minutes, the cells were treated with
The lactate concentration was quantified using a lactate assay kit (Sigma, St. Louis, MO, United States). ASCs were homogenized in 4 × volume of lactate assay buffer. Following centrifugation at 13000 × g for 10 minutes to remove insoluble debris, the supernatant was incubated with 46 mL lactate assay buffer, 2 mL lactate enzyme mix, and 2 mL lactate detection probe at room temperature for 30 minutes. Absorbance was measured at 570 nm using a microplate reader.
ASCs were seeded in 96-well Seahorse plates. Extracellular acidification rate was measured using the Seahorse XFe96 analyzer (Seahorse Biosciences, Billerica, MA, United States) following sequential injections of glucose (10 mmol/L), oligomycin (1 μM), and 2-DG (50 mmol/L). Oxygen consumption rate was analyzed using the same instrument by injecting oligomycin (1 μM), the protonophore uncoupler FCCP (1 μM), and rotenone/antimycin A (0.5 μM each), with final concentrations specified for all reagents.
Total RNA was extracted from ASCs using TRIzol reagent (Invitrogen, Carlsbad, CA, United States). RNA concentration and purity were quantified using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA, United States). First-strand cDNA synthesis and quantitative real-time polymerase chain reaction (qPCR) were performed using the Hifair Advanced One-Step RT-qPCR SYBR Green Kit (Yeasen, Shanghai, China) according to the manufacturer’s instructions. Gene expression levels were analyzed using the 2-ΔΔCt method, with β-actin serving as the internal reference gene.
ASCs were lysed using radioimmunoprecipitation assay lysis buffer. The lysate was collected after centrifugation at 12000 × g for 10 minutes, resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and electrotransferred to polyvinylidene fluoride membranes. Membranes were blocked with 5% non-fat skim milk and incubated with primary antibodies overnight at 4 °C. Following incubation with horseradish peroxidase-conjugated secondary antibodies for 1 hour at room temperature, protein bands were visualized using the Super ECL detection reagent (Yeasen, Shanghai, China).
Mitochondrial staining was performed using MitoTracker Deep Red FM fluorescent dye (MCE, Monmouth Junction, NJ, United States). ASCs were harvested, resuspended in 100 μL of 500 nM MitoTracker Deep Red FM working solution, and incubated at room temperature for 40 minutes. After removal of the staining solution, cells were washed three times with culture medium. Nuclei were counterstained with 4’,6-diamidino-2-phenylindole (MCE, Monmouth Junction, NJ, United States) in mounting medium. Fluorescence was visualized using a fluorescence microscope.
To quantify mitochondrial uptake, ASCs were co-cultured with varying concentrations of mitochondria (mitochondria-to-cell ratios: 0, 125:1, 250:1, 500:1, 1000:1, and 2000:1). Mitochondria were pre-labeled with MitoTracker Green FM dye (MCE, Monmouth Junction, NJ, United States). Briefly, ASCs were incubated with 100 μL of 100 nM MitoTracker Green FM working solution at room temperature for 40 minutes. Following three washes with PBS to remove excess dye, cells were resuspended in flow cytometry buffer. Fluorescence was analyzed using a flow cytometer, and mean fluorescence intensity (MFI) was recorded. Linear regression analysis was performed to correlate MFI with mitochondrial concentration.
Total RNA was extracted using TRIzol reagent according to the manufacturer’s protocol. RNA concentration and purity were assessed using a NanoDrop spectrophotometer. A total of 1 μg of high-quality RNA per sample was used for library construction. Sequencing libraries were generated and pairedend sequencing (2 × 150 bp) was performed on an Illumina NovaSeq 6000 platform (Illumina, CA, United States). Differential expression analysis was performed using the DESeq2 package (v1.38.3) in R (v4.2.0). Genes with an adjusted P-value (false discovery rate) < 0.05 and |log2 fold change| ≥ 1 were considered significantly differentially expressed. RNA sequencing data were analyzed with false discovery rate ≤ 0.05 for multiple testing correction. Differentially expressed RNAs (identified by gene names) were visualized using a heatmap and a volcano plot. Gene set enrichment analysis was performed using the online tool (http://www.broadinstitute.org/gsea) to evaluate the function of the differentially expressed genes. Kyoto Encyclopedia of Genes and Genomes enrichment analysis (https://www.genome.jp/kegg/) was performed to assess the related pathways of the differentially expressed genes.
The enrichment of H3K18 lactylation in the promoter was analyzed using the Pierce magnetic ChIP kit (ThermoFisher, Waltham, MA, United States). Briefly, crosslinking was performed using 1% formaldehyde for 10 minutes, followed by addition of glycine to quench the reaction. Cells were lysed, and the lysate was removed. Cell nuclei were digested with MNase at 37 °C for 15 minutes and sonicated on ice to isolate chromatin. The chromatin sample was incubated with anti-IgG or anti-H3K18 lactylation overnight at 4 °C, followed by incubation with protein A/G magnetic beads for 2 hours at
WT and mutant (MUT) AXIN2, BMPR1B, COL1A1, and OSTN sequences were PCR-amplified and cloned into pGL4 reporter plasmids. These WT or MUT reporter plasmids were transfected into ASCs using Lipofectamine 2000 (Invitrogen, Carlsbad, CA, United States). Forty-eight hours post-transfection, luciferase activities were measured using the dual-luciferase reporter assay system (Promega, Madison, WI, United States). Transcriptional activity was defined as the ratio of firefly to Renilla luciferase activity.
SD rats were randomly divided into five groups [sham, model, model + ASCs, model + ASCs (mt), and model + ASCs (mt) + 2-DG], with six rats per group. The fracture model was generated as previously described[17]. Briefly, rats were anesthetized via isoflurane inhalation and positioned prone. A medial knee incision was made to expose the femoral condyle, followed by insertion of a 1.0 mm diameter Kirschner wire into the femoral marrow cavity and advancement of its tail end into the femoral condyle. The wound was flushed with povidone-iodine solution, and the incision was sutured. Rats were then positioned supine on the fracture induction device, with the supporting platform raised to position the blade adjacent to the thigh. The ramming system was set to a 1.5 mm travel distance (equivalent to a 500 g weight dropped from 35 cm height). The downward force was transmitted to an impact disc, activating the guillotine mechanism to induce a mid-femoral fracture. Buprenorphine HCl was administered for postoperative pain control for three days. The sham group underwent anesthesia and incision only, without Kirschner wire insertion or fracture induction. To investigate the role of ASCs or mitochondria, ASCs (either untreated or mitochondria-stimulated; 1 × 105 cells in 50 μL PBS) were injected into the fracture site one day post-model establishment, administered weekly. Rats in the model + ASCs (mt) + 2-DG group received mitochondria-stimulated ASCs plus 400 mg/kg 2-DG at the fracture site under the same weekly schedule. Eight weeks post-surgery, all rats were euthanized via isoflurane inhalation, and the femurs from the surgical side were harvested.
The fracture callus was fixed in 4% paraformaldehyde for 48 hours, and fracture healing status along with bone microstructural parameters were evaluated using the Skyscan 1176 micro-computed tomography scanner (Bruker, Kontich, Belgium). Three-dimensional reconstruction was performed using NRecon software. Bone microstructural parameters, including bone mineral density, bone volume/total volume, trabecular number, and trabecular thickness, were quantified using the software. Micro-computed tomography analyses were performed by two independent blinded researchers.
The callus tissues at the fracture site were decalcified using 10% EDTA (pH = 7.4) for 21 days. Tissues were then embedded in paraffin and sectioned. After deparaffinization and rehydration, the sections were stained with hematoxylin and eosin using a commercial kit (Solarbio, Beijing, China). Images were captured under a light microscope. Histological analyses were performed by two independent blinded researchers.
Cartilage injury was evaluated using the modified Saffranin O and fast green stain kit (Solarbio, Beijing, China). Paraffin sections were deparaffinized and rehydrated. Subsequently, tissues were stained with Weigert’s hematoxylin solution for 3 minutes, fast green for 5 minutes, and Saffranin O for 5 minutes. After each staining step, sections were washed to remove residual dye. Images were captured under a light microscope. Histological analyses were performed by two independent blinded researchers.
Data were analyzed using GraphPad Prism 8 software. Sample size was chosen based on similar studies and preliminary data to ensure statistical power. n = 3 biological replicates for in vitro experiments; n = 6 rats per group for in vivo experiments. Statistical comparisons were performed using Student’s t-test or one-way ANOVA followed by Tukey’s post hoc test. Results are presented as mean ± SD. Statistical significance was defined as P < 0.05.
In this study, we first isolated ASCs from rats and assessed their multilineage differentiation potential. Following adi
To investigate the effects of platelets on the biological functions of ASCs, the cells were co-cultured with platelets isolated from rat blood. As shown in Figure 2A, platelets did not significantly alter cellular proliferation. Given that dysregulation of glucose metabolism contributes to bone fragility and increased fracture risk[18], we assessed glycolytic activity. Analysis revealed that platelets enhanced glucose uptake (Figure 2B), glycolytic capacity (Figure 2C), mitochondrial respiratory activity (Figure 2D), and lactate production (Figure 2E). Osteogenic differentiation was evaluated by ALP and ARS staining, which showed intensified staining following platelet treatment (Figure 2F and G). Furthermore, platelets upregulated the expression of collagen I, BMP2, and RUNX2 in ASCs (Figure 2H and I). These findings indicate that platelets enhance glycolysis, mitochondrial respiration, and osteogenic differentiation of ASCs.
To confirm that the role of platelets in regulating glycolysis and osteogenic differentiation depends on mitochondrial transfer, we isolated mitochondria from platelets, and platelets depleted of mitochondria were used as the negative control. Osteogenic differentiation was assessed by ALP and ARS staining. The results showed that mitochondria-free platelets did not significantly alter staining intensity, whereas mitochondrial treatment enhanced ALP and ARS signals (Figure 3A and B). Additionally, mitochondrial treatment upregulated the expression of collagen I, BMP2, and RUNX2, while platelets lacking mitochondria had no significant effect (Figure 3C-F). For glycolysis, mitochondrial transfer increased glycolytic capacity (Figure 3G), mitochondrial respiration (Figure 3H), glucose consumption (Figure 3I), and lactate production (Figure 3J). Collectively, these findings demonstrate that platelets promote osteogenic differentiation and glycolysis in ASCs via mitochondrial transfer.
To investigate whether platelet-derived mitochondria are internalized by ASCs, we incubated ASCs with varying concentrations of mitochondria and quantified uptake via flow cytometry. As mitochondrial concentration increased, the proportion of mitochondria-labeled ASCs gradually rose (Figure 4A). Linear regression analysis revealed a strong correlation between mitochondrial concentration and MFI (R2 = 0.998; Figure 4B). Fluorescence imaging confirmed enhanced mitochondrial activity in ASCs following platelet or mitochondrial treatment (Figure 4C), indicating successful transfer of functional mitochondria. Given that mitochondria promoted lactate production (as shown earlier) and lactate can drive histone lactylation, we hypothesized that mitochondrial transfer enhances histone lactylation. Western blot analysis demonstrated that mitochondria increased global lactylation levels and specifically elevated H3K18 lactylation, without affecting H3K27 or H3K9 lactylation (Figure 4D). In addition, we detected whether mitochondria influenced the acetylation of H3K18, and no change of acetylation of H3K18 was found after mitochondria treatment (Supplementary Figure 1). After that, we further screened the writers or erasers of lactylation responsible for this specific modification in the Supplementary Figure 2A and B. We found that P300 might mainly contribute to the lactylation modification of H3K18. Collectively, these results confirm that mitochondria are internalized by ASCs in a dose-dependent manner and selectively promote H3K18 lactylation.
ASCs treated with mitochondria were subjected to RNA-sequencing. Mitochondrial treatment induced significant upregulation and downregulation of multiple RNAs (Figure 5A and B). Gene Set Enrichment Analysis revealed that these differentially expressed RNAs were enriched in pathways associated with DNA replication inhibition, Fanconi anemia, non-alcoholic fatty liver disease, oxidative phosphorylation, and osteogenic differentiation (Figure 5C). Kyoto Ency
Functional rescue experiments confirmed that mitochondrial transfer regulates osteogenesis and glycolysis through H3K18 lactylation. To disrupt lactylation, 2-DG was used to inhibit glycolysis and reduce lactate availability. Mito
Finally, we established a rat fracture model to investigate the role of ASCs, mitochondria, and histone lactylation in vivo by administering ASCs, mitochondria-treated ASCs, or 2-DG (n = 6 per group). Three-dimensional reconstruction images revealed obvious fractures in the model group compared with the sham group. ASCs significantly improved fracture healing, and mitochondria-treated ASCs further enhanced this effect in model rats. Compared with the mitochondria-treated ASCs group, 2-DG exacerbated fracture severity (Figure 7A). The model group exhibited reduced bone mineral density, bone volume/total volume, trabecular number, and trabecular thickness, whereas ASCs reversed these alterations. Mitochondria-treated ASCs further amplified the therapeutic effects of ASCs, while 2-DG counteracted these benefits (Figure 7B). Additionally, ASCs improved histological outcomes and alleviated cartilage damage in fractured rats, with mitochondria-treated ASCs further enhancing these reparative effects; conversely, 2-DG reversed the beneficial impact of mitochondria-treated ASCs (Figure 7C and D). In summary, ASCs facilitate bone repair, and mitochondria further augment this process by promoting histone lactylation.
This study demonstrates that platelets promote glycolysis and osteogenic differentiation of ASCs through mitochondrial transfer. Furthermore, platelet-derived mitochondria induce H3K18 lactylation, which enhances transcription of osteogenesis-related genes. Collectively, this work elucidates the regulatory mechanism by which platelets facilitate bone regeneration via the metabolic reprogramming-epigenetic axis.
MSCs are widely recognized as key cellular mediators in bone tissue engineering due to their multilineage differentiation potential. While numerous studies have focused on bone marrow-derived MSCs for fracture healing and bone defect repair, their clinical application is significantly limited by accessibility constraints, low yield, and donor site morbidity[7]. In contrast, ASCs circumvent these limitations and offer advantages including ease of harvest and robust osteogenic capacity, thereby garnering increasing attention in bone regeneration research. Consequently, we isolated rat ASCs to investigate their osteogenic regulatory mechanisms. Our findings confirm that ASCs promote fracture healing in rats, consistent with prior reports[7,19].
PRP, comprising platelets, growth factors, and cytokines, represents a promising therapeutic strategy for fracture healing[20]. Multiple studies indicate that PRP enhances ASC osteogenic differentiation through diverse molecular pathways, thereby improving bone defects[21-23]. The efficacy of PRP largely hinges on activated platelets, as platelet-derived growth factors stimulate ASC proliferation and osteogenic differentiation[24]. Additionally, platelet lysate maintains ASC osteogenic capacity by modulating chemokine expression[25]. Similarly, we observed that platelets promote ASC osteogenesis, but platelets depleted of mitochondria showed no significant effect, whereas mitochondria isolated from platelets facilitated osteogenic differentiation. This prompted our focus on platelet-derived mitochondria.
During tissue repair, platelet activation triggers mitochondrial fission to meet heightened energy demands by enhancing glycolysis and mitochondrial respiration[26]. This has spurred growing interest in platelet-derived mito
Mitochondrial enhancement of glycolytic flux in recipient ASCs elevated intracellular lactate levels, providing substrates for histone lactylation[29]. We therefore investigated histone lactylation and observed that mitochondrial treatment upregulated H3K18 lactylation. As a direct transcriptional activator in chromatin, histone lactylation[30] prompted us to screen osteogenesis-related factors via RNA-sequencing. We identified four genes whose expression changed post-mitochondrial treatment, all of which were transcriptionally activated by H3K18 lactylation. These findings establish an initial epigenetic mechanism underlying mitochondrial function. Rescue experiments revealed that 2-DG reversed the mitochondrial-mediated promotion of ASC osteogenic differentiation in vitro and bone defect improvement in vivo, confirming that mitochondria exert their effects through H3K18 lactylation. To our knowledge, this is the first systematic investigation of mitochondrial transfer in fracture healing, elucidating its downstream metabolic and epi
However, there are some limitations in the study. Plateletderived mitochondria can be transferred to ASCs through multiple pathways, including tunneling nanotubes, extracellular vesicles/microvesicles, and direct phagocytosis[13]. In our system, the dosedependent uptake and colocalization observed by fluorescence imaging are consistent with an endocytosis/phagocytosis-dependent mechanism. Further investigation using specific inhibitors will be performed in future studies to clarify the dominant route. Moreover, this study demonstrates that platelet-derived mitochondria enhance the therapeutic efficacy of ASCs for fracture healing. The in situ transfer of endogenous platelet-derived mitochondria to ASCs during natural fracture repair will be verified in future studies using transgenic reporter mice. Previous studies have explored its translational applications. For instance, Yao et al’s team[31] has designed an effective hydrogel microneedle patch loaded with stem cell-derived mitochondria-rich extracellular vesicles to gradually release and deliver mitochondria into the wound tissues and boost wound healing. MSCs-derived microvesicles containing functional mitochondria (Mito@euMVs) have been developed for treating mitochondrial dysfunction and aging-related conditions[32]. This study was performed in rats; human cell validation and bone biomechanical testing will be included in future translational research. Time-course studies of mitochondrial transfer and lactylation dynamics will be pursued in future work.
In conclusion, platelet-derived mitochondria promote glycolysis and induce H3K18 lactylation, thereby activating osteogenic differentiation and accelerating fracture healing. These findings not only broaden our understanding of platelet biology but also provide novel insights into the regulatory mechanisms of intercellular mitochondrial transfer. Furthermore, targeting H3K18 lactylation may represent a promising therapeutic strategy for bone regeneration.
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