Engineered Cas9 exosome vesicles as a novel gene editing tool for targeted ASPN editing in osteoarthritis

Abstract

CRISPR-Cas9, an innovative genome-editing technique, holds immense promise in therapeutic applications; nevertheless, the lack of effective delivery methods for in vivo gene editing limits its utility in osteoarthritis (OA) treatment. Recently, exosomes, naturally derived nanosized vesicles secreted by cells, have attracted significant attention as potential vehicles for therapeutic cargo delivery. This study proposes a bioinspired engineered exosome-mediated CRISPR/Cas9 delivery platform for targeted editing of the Asporin (ASPN) gene as a potential precision therapy for OA. Specifically, chondrocyte affinity peptide (Cap)-modified MSC-derived exosomes were employed as natural, biocompatible carriers to deliver CRISPR/Cas9 components specifically to OA-affected chondrocytes, thereby achieving precise and efficient ASPN knockout. Flow cytometry analysis confirmed a modification efficiency of 79.1% for Cap, while the encapsulation efficiency of the ASPN-Cas9 plasmid into exosomes reached 9.5% ± 0.6%. Both in vivo and in vitro investigations revealed that this delivery approach markedly improved cellular uptake and gene-editing efficacy, achieving a substantial reduction of ASPN expression by 61.7%. This, in turn, alleviated ferroptosis, improved mitochondrial function, reduced chondrocyte senescence, inhibited inflammation, and enhanced the cartilage microenvironment. Altogether, these findings strongly suggest the promising therapeutic efficacy of this method in OA models, emphasizing its potential as a precise gene-targeting therapeutic intervention for OA.

Graphical Abstract

Schematic diagram of cartilage-targeted engineered exosomes for specific gene editing in OA Treatment.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12951-026-04158-y.

Background

OA is a prevalent chronic degenerative joint disorder, widely recognized as a major cause of disability worldwide, particularly among older individuals [1]. Typical OA pathological characteristics include the progressive deterioration of cartilage tissue, inflammation of the synovium, and structural alterations within subchondral bone [2]. Disease progression involves a complex interplay among mechanical forces, inflammatory mediators, and genetic susceptibility [3], collectively triggering various pathological pathways, such as ferroptosis induction, intensified lipid peroxidation, mitochondrial dysfunction, impaired redox homeostasis, elevated reactive oxygen species (ROS) production, overexpression of pro-inflammatory cytokines and matrix-degrading enzymes, and increased secretion of senescence-associated secretory phenotype (SASP)-related factors [4]. ASPN, a member of the extracellular matrix (ECM) leucine-rich repeat protein family, was initially identified in human cartilage tissue [5]. Previous investigations revealed ASPN’s interactions with growth factors, notably TGF-β and BMP-2, establishing a regulatory feedback loop that negatively influences chondrogenesis and increases OA vulnerability [6]. Moreover, research by Wang et al. indicated that MiR-4303 acts as a predictive biomarker, reversing lipopolysaccharide (LPS)-induced apoptosis, cell cycle arrest, and reduced cellular viability by targeting ASPN, thus mitigating inflammation in OA chondrocytes [7]. Hence, elevated ASPN expression accelerates cartilage degeneration and intensifies inflammation, reinforcing its crucial role in regulating cartilage homeostasis and highlighting its potential as a therapeutic target for OA treatment.

Gene editing technology involves the precise modification of an organism’s genome, utilizing a range of methods to alter the function of specific genes. Among these approaches, CRISPR-Cas9 has emerged as the most prominent and effective technique, representing a revolutionary advancement in the field of genetic engineering [8]. This groundbreaking method offers significant therapeutic potential for treating diseases such as sickle cell anemia, amyotrophic lateral sclerosis (ALS), cancer, and cystic fibrosis [9, 10]. By delivering exceptional precision in genome editing, it opens new avenues for advancements in personalized medicine, innovative disease therapies, and the overall development of biotechnology. Although CRISPR technology holds transformative potential, delivering it safely and efficiently to target cells continues to pose significant challenges due to biological barriers and the constraints of tissue-specific delivery systems. At present, viral vectors like lentivirus and adeno-associated virus (AAV) are commonly used for in vivo delivery [11, 12]. However, these methods are not without limitations, including immune system activation and restricted packaging capacity [13]. Exosomes (50–150 nm), small vesicles secreted by cells, naturally carry genetic material and serve as essential mediators of intercellular communication. Recently, they have attracted significant interest as potential vehicles for gene delivery [14]. In contrast, engineered exosomes, an advanced form of non-viral nanocarrier technology, modified with chondrocyte-affinity peptides (Cap), have shown exceptional promise in OA treatment [15]. These exosomes provide distinct benefits, including superior cargo capacity, natural compatibility with biological systems, low immunogenicity, and highly specific tissue-targeting capabilities [16]. Consequently, the development of CRISPR/Cas9-integrated engineered exosomes, tailored for chondrocyte-specific targeting to genetically alter the ASPN gene in OA chondrocytes, represents a revolutionary advancement in OA treatment.

This study presents an innovative strategy to address the progression of OA. As illustrated in Fig. 1, we initiated the process by incorporating ASPN-Cas9 into exosomes derived from mesenchymal stem cells (MSCs), resulting in the formation of Exo/ASPN-Cas9. To increase targeting specificity, these exosomes were further engineered with the lysosome-associated membrane protein 2b (Lamp2b) gene and modified with a Cap, leading to the creation of Cap-Exo/ASPN-Cas9. The engineered exosomes were comprehensively characterized, with their effects assessed both in vitro and in vivo. Through transcriptomic sequencing, we also identified and validated their potential molecular mechanisms. By targeting genetic factors involved in cartilage degradation, this approach shows great potential for delivering more precise and effective OA therapies.

Fig. 1.

Schematic diagram of cartilage-targeted engineered exosomes for specific gene editing in OA Treatment

Materials and methods

Cell culture

Chondrocytes were isolated from cartilage tissues harvested from the knee joints of 3–5-day-old C57BL/6 mice. The cartilage samples underwent enzymatic digestion at 37 °C for 6 h with collagenase type II (0.2%; Sigma-Aldrich, USA). Following digestion, the resulting cell suspension was subjected to centrifugation at 1500 rpm for 5 min. The pelleted cells were then resuspended and subsequently cultured at 37 °C in a humidified incubator containing 5% CO₂, using DMEM/F-12 (Gibco, USA) supplemented with fetal bovine serum (FBS, 10%; Gibco, USA) and penicillin-streptomycin (PS, 1%). For downstream experiments, only chondrocytes that reached passage two or higher were selected. Bone marrow-derived mesenchymal stem cells (MSCs) were extracted aseptically from the femurs of 4–6-week-old C57BL/6 mice and cultivated in α-MEM medium (Gibco, USA) containing 10% FBS and 1% PS. RAW264.7 cells were propagated using DMEM with identical concentrations of FBS (10%) and PS (1%), following the same standard conditions established for MSCs and chondrocytes.

Animals

Eight-week-old male C57BL/6 mice obtained from the Chinese Academy of Sciences’ Institute of Zoology were used in this study. All experimental procedures involving animals were authorized by the Animal Care and Use Committee at Wenzhou Medical University (approval number: wydw2025-0032). Osteoarthritis was experimentally induced through surgical destabilization of the medial meniscus (DMM), a previously validated method causing joint degeneration [44]. After surgery, mice were randomly distributed into seven groups and subjected to weekly intra-articular administrations (10 µL each) of Exo, Cap-Exo, Exo/ASPN-Cas9, Cap-Exo/ASPN-Cas9, celecoxib, or a corresponding volume of PBS for four weeks consecutively. To generate OA models with different severity, the duration of DMM-induced degeneration was varied. Following treatment, joint tissues were collected and analyzed histologically, molecularly, and through imaging methods.

Bioinformatic analysis

Gene expression data from healthy and osteoarthritic cartilage tissues were acquired from the Gene Expression Omnibus (GEO) repository, comprising datasets GSE114007, GSE169077, and GSE207881. Differentially expressed genes (DEGs) relevant to OA were extracted from raw datasets. Prior to DEG identification, data underwent normalization and batch-effect adjustment. The limma package in R software was employed for analysis, with DEGs selected based on stringent statistical thresholds: adjusted p-value < 0.05 and |log2 fold change| > 1. Visualization of DEGs included heatmaps and volcano plots generated with R packages heatmap and ggplot2, respectively. Furthermore, shared DEGs across datasets were identified and visualized via Venn diagrams constructed using R.

Plasmid construction

CRISPR-Cas9 plasmids for gene editing were constructed utilizing the pSpCas9 backbone from Addgene (USA). An single-guide RNA (sgRNA) targeting the ASPN gene was designed with an online CRISPR design platform and cloned into the pSpCas9 plasmid at the BbsI restriction enzyme site. The sgRNA sequence targeting ASPN is depicted in Figure S1A. The FLAG-Lamp2b plasmid was commercially obtained from Shanghai Newup Biotech Co., Ltd., and its plasmid structure is provided in Figure S1B.

Exosome isolation and characterization

Exosome purification was performed by differential ultracentrifugation from MSC culture supernatants. Initially, conditioned media were centrifuged at 300 × g and subsequently at 2000 × g, each for 10 min, removing cellular material and debris, followed by a 10,000 × g spin for 30 min to discard apoptotic fragments. Exosomes were ultimately collected by ultracentrifugation at 100,000 × g for 2 h at 4 °C, after which the pellets were stored at − 80 °C. MSCs were transiently transfected with Cap-FLAG-Lamp2b plasmids encoding the membrane protein Lamp2b fused to a FLAG tag at a concentration of 4 µg/well, as optimized by preliminary tests. Subsequently, exosomes presenting Cap-FLAG-Lamp2b (Cap-Exo) were collected. For encapsulation of the CRISPR-Cas9 system, exosomes were loaded with ASPN-Cas9 plasmids via electroporation. Specifically, 20 µg ASPN-Cas9 plasmid and 100 µg Cap-Exo (based on total protein content) were combined in 400 µL electroporation buffer (1 mM MgCl₂, 10 mM K₂HPO₄/KH₂PO₄, pH 7.2). The mixture was transferred to a 0.4 cm electroporation cuvette and square wave pulses were applied at 200 V and 150 µF using the Gene Pulser Xcell™ system (Bio-Rad) [45]. Exosome morphology was evaluated via Transmission Electron Microscopy (TEM, Hitachi, Japan). Briefly, exosome samples were fixed with 2.5% glutaraldehyde, dehydrated, and then imaged using TEM. Western blot (WB) analysis was employed to identify exosome markers, including positive markers TSG101, CD63, and CD81 (Abcam, USA), as well as the negative marker Calnexin. Nano-flow cytometry (NanoFCM, Xiamen Fuliu Biotechnology Co., Ltd.) was employed to evaluate the concentration and particle size distribution of the samples for high-resolution detection.

Cellular uptake assay

For the cell uptake study, red fluorescent TRITC-conjugated phalloidin (Solarbio, China) was used to label the cytoskeleton, while green fluorescent PKH67 dye (Sigma-Aldrich, USA) was employed to label exosomes. DAPI staining (Beyotime, China) was used for visualizing cell nuclei. Chondrocytes were treated overnight with exosomes labeled with the fluorescent dye PKH67. Following fixation, the internalization of labeled exosomes by chondrocytes was visualized via laser scanning confocal microscopy (Leica, Germany).

T7E1 assays

The T7E1 assay was conducted to evaluate the efficiency of CRISPR-Cas9-mediated gene editing.Genomic DNA was isolated from treated chondrocytes, and the ASPN target region was amplified via PCR with specific flanking primers. The amplified DNA underwent denaturation and reannealing, then was digested with T7 Endonuclease I (New England Biolabs, USA). The cleavage products were separated through gel electrophoresis to confirm the presence of insertions/deletions (Indels), and the results were analyzed using ImageJ [46]. The Indel percentage was calculated as follows:

Indel percentage = 100 × (1 − (1 − cleavage fraction) 1/2), where the cleavage fraction = band intensity of each cleaved band / (band intensity of each cleaved band + band intensity of the uncut band).

Cell viability assay

The viability of chondrocytes was assessed by employing a Cell Counting Kit-8 assay (Dojindo, Japan). Briefly, cells (5 × 10⁴ per well) were plated onto 96-well plates, cultured, and exposed for 24, 48, or 72 h to Exo, Exo/ASPN-Cas9, or different concentrations of Cap-Exo/ASPN-Cas9. Following incubation, each well received an addition of 10 µL CCK-8 reagent and was incubated for another 2 h at 37 °C. The absorbance at a wavelength of 450 nm was subsequently measured utilizing a microplate spectrophotometer (Thermo Fisher Scientific).

Western blot

Cellular proteins were extracted utilizing RIPA buffer (Beyotime, China), and protein concentrations were quantified through a BCA protein assay (Biyuntian, China). Subsequently, equal amounts of protein were electrophoretically resolved by SDS-PAGE, transferred onto PVDF membranes (Millipore, USA), and blocked at 37 °C with 5% bovine serum albumin (BSA, Sigma, USA) for 2 h. The membranes were then incubated overnight (4 °C) with specific primary antibodies listed in Table S1, followed by incubation with corresponding HRP-conjugated secondary antibodies for 2 h. Protein signals were detected using an enhanced chemiluminescence (ECL) reagent.

Flow cytometry analysis

RAW264.7 cells were seeded into 6-well plates and grown to approximately 80–90% confluence. Cells were then stimulated for 24 h with lipopolysaccharide (LPS, 100 ng/mL) combined with interferon-γ (IFN-γ, 20 ng/mL) to induce M1 macrophage polarization. After stimulation, cells were harvested and labeled at 37 °C for 30 min using anti-F4/80-FITC and anti-CD86-PE antibodies (Abcam, USA). Following washing and resuspension in PBS, flow cytometric analysis was carried out on a BD FACSCalibur cytometer (USA).

Quantitative real-time PCR

RNA was extracted from cells with TRIzol reagent (Thermo Fisher Scientific, USA), followed by reverse transcription into cDNA using the PrimeScript RT reagent kit (Takara Bio, Japan). Quantitative PCR analysis was performed utilizing the StepOnePlus Real-Time PCR System along with SYBR Green PCR Master Mix (Applied Biosystems, USA). Relative gene expression levels were normalized against GAPDH and calculated according to the 2^−ΔΔCt method. The primer sequences used are presented in Table S2.

ELISA

Inflammatory markers, including TNF-α, IL-18, PGE2, and nitrite levels, were quantified from collected cell culture supernatants using commercially available ELISA kits and Griess reagent (eBioscience, USA), strictly following the manufacturer’s guidelines. The absorbance at 450 nm was subsequently measured by a microplate reader (Thermo Fisher Scientific, USA).

Immunofluorescence (IF) analysis

Cells cultured in 24-well plates underwent fixation with 4% paraformaldehyde and permeabilization utilizing 0.3% Triton X-100, followed by blocking with 5% BSA. Samples were then incubated overnight at 4 °C with primary antibodies targeting Collagen-II, COX2, MMP-13, SLC7A11, GPX4, or Nrf2, followed by a 1-hour incubation with AlexaFluor^® 488 or AlexaFluor^® 594 secondary antibodies. Nuclei were visualized via DAPI staining. Fluorescent images were obtained and analyzed using an inverted fluorescence microscope (Zeiss, Germany).

Cell toluidine blue (TB) and Safranin O (SO) staining

Chondrocyte cultures or cartilage tissue sections were stained with TB and SO (Solarbio, China) as per manufacturer’s instructions to visualize cartilage matrix integrity and proteoglycan content. Samples were evaluated using a light microscope (Olympus, Japan).

Transcriptome sequencing

Transcriptomic alterations in chondrocytes following Cap-Exo/ASPN-Cas9 treatment relative to untreated control cells were profiled by RNA sequencing. TRIzol reagent was utilized to isolate total RNA, whose integrity was verified by the Agilent 2100 Bioanalyzer and quantified via Nanodrop spectrophotometry, ensuring RIN scores exceeded 7.0. Sequencing libraries were constructed using Illumina’s TruSeq RNA library kit, and subsequently sequenced on the HiSeq 2500 instrument (Illumina, USA) generating 150 bp paired-end reads. Cutadapt software (v1.9) was employed to filter out adapter sequences and low-quality reads from the raw data. Differentially expressed genes (DEGs) were defined based on stringent criteria (adjusted p-value < 0.05, fold-change > 1.5), and results were visualized through volcano plots and heatmaps to depict gene expression profiles. Furthermore, functional enrichment analyses, including Gene Ontology (GO), Kyoto Encyclopedia of Genes and Genomes (KEGG), and Gene Set Enrichment Analysis (GSEA, v4.1.0), were performed to identify significantly enriched biological pathways (p < 0.05, False Discovery Rate (FDR) < 0.25) affected by Cap-Exo/ASPN-Cas9 intervention.

Molecular docking

Three-dimensional structural data of ASPN and Nrf2 were retrieved from the Protein Data Bank (PDB). Molecular docking analysis between ASPN and Nrf2 was performed using AutoDock Vina, and the interaction interfaces and binding affinities were visualized using PyMOL software (version 2.2.0).

Immunoprecipitation

To evaluate protein-protein interactions, cell lysates were incubated overnight at 4 °C with antibodies targeting ASPN or Nrf2; normal rabbit IgG served as the negative control. Subsequently, the immune complexes were incubated for 2 h at 4 °C with Protein A/G beads (Thermo Fisher Scientific, USA). The precipitated complexes were washed thoroughly and subjected to WB analysis.

Detection of intracellular ROS, iron and lipid-ROS

Fluorescent probes DHE and DCFH-DA (Invitrogen, USA) were used to measure intracellular reactive oxygen species (ROS) generation. Chondrocytes were stained with diluted probes in the dark for 20 min at 37 °C, subsequently rinsed in PBS, and visualized using confocal microscopy. The intracellular concentration of Fe²⁺ was quantified using an iron-specific fluorescence-based probe (Dojindo, Japan) according to the manufacturer’s instructions. Additionally, the lipid peroxidation level was evaluated via BODIPY-C11 staining (Invitrogen, USA), and fluorescence intensities were quantitatively assessed using ImageJ software.

TEM

Chondrocyte samples were fixed at 4 °C in 2.5% glutaraldehyde solution for 2 h and subsequently incubated in 1% osmium tetroxide for 1 h. After fixation, the samples underwent graded ethanol dehydration and were embedded in Epon resin. Thin sections approximately 70 nm thick were sliced using an ultramicrotome, placed onto copper grids, and subsequently stained with uranyl acetate and lead citrate to enhance electron microscopic contrast. Mitochondrial morphology, including size, cristae structure, and membrane integrity, was examined using JEM-1200EX (JEOL, Japan). Images were captured to assess mitochondrial damage, including swelling, membrane rupture, and cristae loss, as indicators of dysfunction.

Mitochondrial function assays

Mitochondrial oxidative stress was evaluated by detecting mitochondrial superoxide generation with MitoSox Red dye (Invitrogen, USA). To measure mitochondrial superoxide production, chondrocytes underwent staining at 37 °C with 5 µM MitoSox Red dye for 30 min, after which fluorescence signals were captured by fluorescence microscopy. Furthermore, mitochondrial mass and integrity were determined using MitoTracker Red (Invitrogen, USA); cells were incubated for 30 min at 37 °C in medium containing 200 nM of the dye, and fluorescence microscopy images were subsequently analyzed. Additionally, mitochondrial membrane potential (ΔΨm) was measured using JC-1 staining (Thermo Fisher Scientific, USA; 5 µg/mL, 20 min at 37 °C) [47].

SA-β-Gal staining

For cellular senescence assessment, chondrocytes were fixed and subjected to staining with SA-β-Gal solution adjusted to pH 6.0 according to manufacturer’s protocol. Senescent cells, identified by blue staining, were counted, and the percentage of SA-β-Gal-positive cells was calculated to determine cellular senescence rates.

Radiographic and microcomputed tomography scans (Micro-CT)

Eight weeks post-surgery, mice were anesthetized, and X-ray imaging of the knee joints was performed using a radiographic system (Faxitron, USA). X-ray images were captured to visualize skeletal abnormalities, such as osteophyte formation and joint space narrowing, which are characteristic of OA. Moreover, knee joints were scanned using micro-computed tomography (microCT, Bruker, Germany) at a resolution of 9 μm to obtain high-resolution three-dimensional images. MicroCT image reconstruction was performed using NRecon software (Bruker, Germany), facilitating visualization of subchondral bone remodeling, joint space narrowing, and osteophyte formation. Quantitative measurements of joint space width and osteophyte dimensions were performed using CTAn software (Bruker, Germany), allowing precise evaluation of bone density and structural alterations within the joint.

Histopathology

The knee joint tissues were subjected to fixation in 10% formalin, decalcification using EDTA solution, paraffin embedding, and subsequent sectioning for histopathological evaluation. Cartilage morphology and tissue integrity were assessed by histochemical staining, including Safranin O/Fast Green, toluidine blue (TB), and hematoxylin-eosin (H&E). The Osteoarthritis Research Society International (OARSI) scoring method was employed to quantitatively evaluate cartilage deterioration, while synovial inflammation severity was scored using an established synovitis scoring protocol [48, 49].

Immunohistochemcal (IHC) analysis

Expression levels of iNOS, COX2, ACLS4, MMP-13, Collagen II, and Nrf2 in knee joint sections were assessed by immunohistochemistry. Following conventional deparaffinization, hydration, and antigen-retrieval processes, tissues were incubated at 4 °C overnight with the corresponding primary antibodies. HRP-linked secondary antibodies were then added, and sections incubated at ambient temperature for 1 h. Protein signals were visualized and digitally scanned using a slide imaging system (Affinity, China), and staining intensities were quantitatively assessed via ImageJ software.

Statistical analysis

Data are presented as mean ± standard deviation (SD). Statistical analyses were carried out with GraphPad Prism 8.0 software. Comparisons between two independent groups with normally distributed data were performed using an independent two-tailed Student’s t-test. For multiple-group comparisons, one-way analysis of variance (ANOVA) was conducted, followed by Tukey’s post hoc tests. A value of p < 0.05 was considered statistically significant.

Results

ASPN is a critical biomarker involved in the pathological progression of OA

Gene expression data from GEO datasets GSE114007, GSE169077, and GSE207881 were analyzed using R-based bioinformatics tools to identify DEGs associated with OA. These DEGs, associated with the pathogenesis of OA, were identified through rigorous analysis and are visually represented using volcano plots and heatmaps (Fig. 2A, B). To identify commonalities across the datasets, a Venn diagram was constructed, which highlighted the overlap of DEGs, revealing 20 DEGs that were consistently upregulated in OA across the three datasets (Fig. 2C).

Fig. 2.

ASPN expression is positively correlated with OA. (A) Volcano plots of DEGs from three OA-related GEO databases. (B) Heatmap of DEGs from three OA-related GEO databases. (C) The Venn diagram showed the overlap between all highly expressed DEGs in these three databases. (D) The expression levels of 20 overlapping highly expressed DEGs in human cartilage were ranked from high to low. (E) Expression of ASPN in the above three databases. (F) Representative images of histological staining of mouse knee joint sections and immunohistochemical assay of ASPN in cartilage. (G) OARSI scoring of mouse knee cartilage. (H) Quantification of positive areas of ASPN expression in cartilage. (I) WB analysis was used to detect the expression of ASPN in chondrocytes of different degrees of OA. (J) Correlation analysis between ASPN positive expression and cartilage OARSI score. (ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001; n = 3)

Among the 20 upregulated DEGs, ASPN was identified as the gene exhibiting the most significant differential expression between OA and normal groups, as shown in Fig. 2D. This gene consistently demonstrated marked upregulation across all three datasets, further confirming the robustness of its association with OA (Fig. 2E). The presence of ASPN in OA was further validated in histological analysis. In Fig. 2F, histological staining of knee joint tissues from OA mouse models of varying severity was performed. As expected, the severity of OA correlated with increased roughness of the knee joint surface, more pronounced cartilage erosion, and higher pathological scores for OA (Fig. 2G). These observations underscore the progressive nature of OA in the animal model. Interestingly, in parallel with the worsening OA severity, the positive expression of ASPN was increasingly evident in the affected tissues (Fig. 2H), strongly suggesting a close relationship between ASPN expression and the progression of OA. To further investigate this correlation, we collected chondrocytes from OA models representing different stages of severity and proceeded with protein extraction. ASPN expression levels were quantified using WB analysis. The findings demonstrated a progressive rise in ASPN expression correlating with OA severity (Fig. 2I). A robust positive association between elevated ASPN levels and OA severity (Fig. 2J) emphasizes the critical role of ASPN in the pathogenesis of OA, further highlighting its promise as an indicator of disease progression.

Preparation and characterization of chondrocyte-targeting engineered exosomes

Considering ASPN’s identification as an essential marker linked to OA progression, targeted deletion of the ASPN gene may offer therapeutic benefits. Thus, we established a CRISPR/Cas9-mediated gene-editing platform targeting ASPN, termed ASPN-Cas9. The ASPN-Cas9 system employs a single-guide RNA (sgRNA) designed to precisely guide Cas9 nuclease to the ASPN genomic site, enabling accurate gene cleavage (Fig. 3A). The pSpCas9 plasmid was engineered to express sgRNA sequences specific for ASPN. Considering that ASPN is highly abundant in chondrocytes, we developed exosomes designed specifically for chondrocyte-targeting by modifying their surfaces with a peptide possessing affinity for chondrocytes (Cap), fused to lysosomal-associated membrane protein 2b (Lamp2b). These engineered exosomes were subsequently named Cap-Exo (Fig. 3B). Subsequently, ASPN-Cas9 plasmids were encapsulated within Cap-Exo via optimized electroporation, yielding Cap-Exo/ASPN-Cas9 complexes.

Fig. 3.

Extraction and characterization of cartilage-targeted exosomes. (A) Schematic diagram of the design of the ASPN-Cas9 system. (B) Schematic diagram of ASPN-sgRNA sequence and cartilage-targeted exosome generation. (C) TEM images of Exo, Exo/ASPN-Cas9 and Cap-Exo/ASPN-Cas9. (D) Detection of exosome marker proteins by WB. (E) Concentration and particle size of Exo, Exo/ASPN-Cas9 and Cap-Exo/ASPN-Cas9 detected by nanoflow cytometry. (F) Stability test of exosomes at 4 °C for 7 days. (G) Representative images of intracellular uptake in chondrocytes. (H) Frequency of ASPN indel mutations detected in chondrocytes by T7E1 assay. (I) Gene expression of ASPN in chondrocytes after treatment with different groups. (J, K) WB detection of ASPN protein expression in chondrocytes after treatment in different groups. (n = 3, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, *p < 0.05; **p < 0.01; ***p < 0.001)

Morphologically, Cap-Exo/ASPN-Cas9 appeared as oval-shaped spheres with a diameter of approximately 80–100 nm, similar to conventional exosomes (Fig. 3C). Next, we tested the surface exosomal markers of Cap-Exo/ASPN-Cas9. The results were similar to those of conventional Exo, and WB confirmed that Cap was successfully incorporated onto the surface (Fig. 3D, S2). Nanoflow cytometry confirmed that the average particle size of Cap-Exo/ASPN-Cas9 was 79.89 nm and the concentration was 1.32E + 10 particles/mL. These parameters were similar to those of traditional Exo and Exo/ASPN-Cas9. However, due to the positive charge of Cap, its Zeta potential was 1.7 mV, which was more conducive to binding to negatively charged cartilage (Fig. 3E). In addition, all three types of exosomes showed good stability, maintaining a consistent size at least for 7 days at 4 °C (Fig. 3F). To determine whether our chondrocyte-targeted engineered exosomes could more efficiently enter chondrocytes, we labeled the exosomes with PKH67. The results showed that engineered exosomes carrying the Cap sequence exhibited significantly higher signal intensity compared to exosomes without surface-functionalized Cap sequences (Fig. 3G, S3). To quantitatively determine the modification rate of exosome surface Cap, we labeled CAP with Cy5 and used flow cytometry to evaluate the modification efficiency. We determined that when 4 µg of plasmid was used per well (6-well plate format), the optimal Cap modification rate reached a steady state, with a surface modification efficiency of approximately 79.1% (Figure S4A, B). To assess encapsulation efficiency, we employed the Quant-iT™ PicoGreen dsDNA Assay Kit (Invitrogen). Serial dilutions of ASPN-Cas9 plasmid DNA (ranging from 0 to 200 ng/mL) were prepared to establish a double-stranded DNA standard curve, and fluorescence intensities were recorded at excitation/emission wavelengths of 480/520 nm. As illustrated in Figure S4C, the linear correlation coefficient (R² = 0.998) indicated high reliability of the assay. Loading efficiency was determined as the proportion of encapsulated plasmid DNA relative to the total DNA amount initially loaded. As shown in Figure S4D, our results showed a loading efficiency of 9.5% ± 0.6%, which is consistent with the previously reported efficiency of loading plasmids into exosomes by electroporation, especially for large plasmids (> 9 kb), which confirms the feasibility of our delivery strategy. We then investigated the potential of exosome-mediated ASPN-Cas9 to disrupt the ASPN gene in vitro. As shown in Fig. 3H, ASPN-Cas9 achieved a noticeable indel frequency at different loci, indicating that ASPN-Cas9 facilitated efficient gene editing at these loci. We then used qRT-PCR to detect the expression level of ASPN in chondrocytes. Cap-Exo/ASPN-Cas9 showed the most effective inhibitory effect on ASPN expression, with ASPN expression significantly reduced by 67.3% (Fig. 3I). Further confirmation by WB analysis demonstrated effective ASPN suppression via Cap-Exo/ASPN-Cas9. Specifically, the protein level of ASPN in cells treated with Cap-Exo/ASPN-Cas9 decreased to 0.37 compared with the Exo-treated group. In contrast, cells treated with Exo/ASPN-Cas9 exhibited ASPN protein levels at 0.58 relative to the control, aligning with our prior findings and validating the high gene-editing efficiency of the chondrocyte-targeted engineered exosomes (Fig. 3J, K).

To demonstrate the targeting of Cap-expressing exosomes to chondrocytes in vivo, we performed small animal fluorescence imaging experiments. Additionally, both CAP-Exo/ASPN-Cas9 and Exo/ASPN-Cas9 complexes were fluorescently labeled using DiR, a near-infrared dye. A volume of 10 µL labeled exosomes was injected intra-articularly into mouse knee joints. In vivo fluorescence imaging performed at intervals of 4, 24, 48, and 72 h post-administration revealed significantly enhanced fluorescence intensity and retention in the CAP-Exo/ASPN-Cas9-treated group compared to controls. After 72 h, the control group had almost no retention in the knee joint, while CAP-Exo/ASPN-Cas9 still had a large amount of accumulation in the knee joint, indicating that its retention and targeting in the cartilage area were enhanced (Figure S5A, B).

To evaluate the articular cartilage penetration ability and cellular uptake efficiency of both CAP-Exo/ASPN-Cas9 and Exo/ASPN-Cas9, a volume of 10 µL DiR-labeled exosomes was injected intra-articularly into mouse knee joints. After 48 h, the mouse cartilages were cryo-sectioned and observed by laser confocal microscopy. As shown in Figure S16A, Exo/ASPN-Cas9 revealed a very limited penetration ability. Significantly, CAP-Exo/ASPN-Cas9 had distributed throughout the cartilage after penetration for 48 h. Meanwhile, we analyzed the uptake efficiency of cartilage to exosomes, 56.00% ± 3.60% CAP-Exo/ASPN-Cas9 were taken up and 18.23% ± 3.12% Exo/ASPN-Cas9 were taken up in mouse cartilage after 48 h (Figure S16B). These results convincingly verified that CAP-Exo/ASPN-Cas9 are capable of penetrating into the deep zone of the articular cartilage and exhibit improved cartilage penetration ability.

To further validate the potential for in vivo and vitro application of Cap-Exo/ASPN-Cas9, we performed toxicity and cell proliferation assays. In vitro, Exo, Exo/ASPN-Cas9, and Cap-Exo/ASPN-Cas9 all showed a positive effect on promoting chondrocyte proliferation. Notably, when the concentration of Cap-Exo/ASPN-Cas9 was 10 µg/mL, the effect was significantly enhanced (Figure S6). Therefore, this concentration was selected for use in the following in vitro experiments.

Cap-Exo/ASPN-Cas9 exhibits enhanced anti-inflammatory effects in vitro

One of the key hallmarks of OA is local joint and synovial inflammation. Chronic low-grade inflammation in OA has long been recognized as a major driving force behind joint degradation and the progression of disease. An interesting aspect of OA progression is the exacerbating role of M1 macrophage-driven synovitis, which contributes to the inflammatory milieu within the joint [17]. We utilized flow cytometry to evaluate M1 macrophage proportions across various treatment groups, aiming to elucidate the impact of M1 macrophage dysregulation in OA. The results showed that the proportion of M1 increased significantly after LPS/IFN-γ induction, reaching 73.24%. The use of Exo had a certain inhibitory effect on M1, with a proportion of about 58.54%. Cap-Exo/ASPN-Cas9 and Exo/ASPN-Cas9 embedded with ASPN-Cas9 both showed significantly stronger inhibitory effects on M1 macrophages than conventional Exo, with the proportions of M1 being 49.36% and 46.61%, respectively. This suggests that knocking out ASPN effectively inhibit the polarization of M1 macrophages, thereby reducing the inflammatory response. No statistically significant difference was observed between Cap-Exo/ASPN-Cas9 and Exo/ASPN-Cas9, potentially due to the targeted nature of the exosomes (Fig. 4A, B).

Fig. 4.

Cap-Exo/ASPN-Cas9 has enhanced anti-inflammatory and ECM maintenance effects. (A, B) RAW264.7 cells were used for flow cytometry to detect the proportion of M1 macrophages after different treatments. (C, D) WB was used to detect the expression levels of inflammatory proteins related to primary chondrocytes after different treatments. (E) WB was used to detect the expression levels of ECM-related proteins. (F) Representative images of TB and SO staining of chondrocytes after treatment in different groups. (G, H) Representative IF images of the expression of Collagen-II in chondrocytes after treatment in different groups. (I, J) Representative IF images of MMP-13 expression in chondrocytes after treatment in different groups. (n = 3, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001)

To explore the potential anti-inflammatory effect of Cap-Exo/ASPN-Cas9, we used ELISA to measure the levels of several key inflammatory mediators in the supernatant of primary chondrocytes, including TNF-α, PGE2, IL-18, and Nitrite. The results clearly demonstrated that Cap-Exo/ASPN-Cas9 exerted a highly effective inhibition of these inflammatory mediators compared to conventional exosomes (Exo) or Exo/ASPN-Cas9 lacking the Cap (Figure S7A). Further WB analysis confirmed the effective inhibitory role of Cap-Exo/ASPN-Cas9 on the expression of crucial inflammatory proteins. As illustrated in Figs. 4C and D, exposure to TBHP substantially elevated the levels of inflammatory markers such as INOS, COX2, and TNF-α, approximately two- to three-fold compared to the control group. However, treatment with Cap-Exo/ASPN-Cas9 significantly attenuated the expression of these proteins relative to Exo and Exo/ASPN-Cas9 treatments. Furthermore, qPCR results indicated that although TBHP markedly enhanced inflammatory gene expression, Exo alone provided only minimal anti-inflammatory effects. Exo/ASPN-Cas9 could play a better role in inhibiting inflammation than Exo, but not as strong as Cap-Exo/ASPN-Cas9 (Figure S7C). Further experiments again supported the pronounced anti-inflammatory capacity of chondrocyte-targeted engineered exosomes. To more precisely demonstrate Cap-Exo/ASPN-Cas9’s inflammation-suppressing potential, IF staining was applied to measure inflammation-associated proteins. Results indicated a remarkable elevation of the inflammatory mediator COX2 following TBHP stimulation, reflected by increased cytoplasmic fluorescence intensity reaching approximately 2.31-fold greater levels relative to untreated controls. The use of Exo and Exo/ASPN-Cas9 also reduced the expression of COX2 protein, with fluorescence intensity of about 1.88 and 1.43 times that of the control group. It is worth noting that compared with the first two groups, Cap-Exo/ASPN-Cas9 treatment significantly reduced the expression of COX2, with a fluorescence intensity of 1.22 times that of the control group, highlighting its excellent anti-inflammatory effect (Figure S7B, D). The anti-inflammatory properties of Cap-Exo/ASPN-Cas9 appear to be pivotal in regulating inflammation and decelerating the pathological progression of OA.

Cap-Exo/ASPN-Cas9 suppresses ECM degradation to maintain the chondrocyte microenvironment in vitro

The ECM is a complex network structure located outside the cells, composed of various molecules, including proteins, glycosaminoglycans, and glycoprotein [18, 19]. It offers physical support to cells and is essential for biological processes like cell growth, migration, differentiation, signal transduction, and tissue repair. Enhanced degradation of joint cartilage ECM significantly contributes to the development and advancement of OA. To evaluate the ability of Cap-Exo/ASPN-Cas9 to maintain the chondrocyte microenvironment, we first visualized the effect of Cap-Exo/ASPN-Cas9 on ECM in vitro. WB analysis revealed that compared to Exo and Exo/ASPN-Cas9, Cap-Exo/ASPN-Cas9 treatment significantly reduced the expression of key ECM-degrading components, such as ADAMTS4 and MMP-13, while restoring the levels of Aggrecan and Collagen-II (Fig. 4E, S7E). This remarkable ability to maintain the chondrocyte ECM may be related to its cartilage-targeting specificity. Additionally, qPCR results confirmed that Cap-Exo/ASPN-Cas9 effectively downregulated the expression of MMP-13 and other ECM-degrading enzymes, including ADAMTS4, compared to the control group (Figure S7F). Analysis of collagen II and aggrecan, two major structural components of cartilage, by TB staining and safranin staining further confirmed the reduction of ECM degradation by Cap-Exo/ASPN-Cas9 (Fig. 4F), which is consistent with our previous findings. IF results showed that TBHP could induce the destruction of chondrocyte ECM, such as the degradation of ECM component collagen II and the increased synthesis of MMP-13, which is a key enzyme involved in ECM degradation. The IF of collagen II in the TBHP group decreased by about 0.28 times that of the control group (Fig. 4G, H), while the fluorescence expression intensity of MMP-13 was about 2.67 times that of the control group (Fig. 4I, J). Although the use of Exo and Exo/ASPN-Cas9 alleviated this destruction to a certain extent, it was still insufficient compared with Cap-Exo/ASPN-Cas9. Moreover, Cap-Exo/ASPN-Cas9 significantly counteracted TBHP-induced degradation of collagen II and reduced MMP-13 expression. The fluorescence intensity of collagen II was restored to around 0.75-fold compared to controls, while MMP-13 fluorescence was reduced to approximately 1.30-fold, indicating the maintenance of ECM homeostasis. In conclusion, these findings suggest that Cap-Exo/ASPN-Cas9 effectively suppresses ECM degradation and helps maintain the chondrocyte microenvironment by downregulating key ECM-degrading enzymes. From a superficial perspective, this provides a promising strategy for protecting cartilage integrity in OA.

CRISPR-Cas9-mediated ASPN knockout May exert its effect by interacting with the Nrf2 transcription factor

Preliminary data suggest that targeted ASPN disruption via CRISPR-Cas9 is efficacious. To uncover the molecular pathways involved in the ASPN knockout mechanism, high-throughput sequencing and bioinformatics analysis were implemented. Compared to controls, cells treated with Cap-Exo/ASPN-Cas9 exhibited 2,265 DEGs, with 1,075 genes upregulated and 1,190 genes downregulated. The DEGs were visually represented through heatmaps and volcano plots (Fig. 5A, B). Interestingly, we found that NFE2L2 was one of the upregulated DEGs among the 1,075 genes, and through molecular docking, we investigated the interaction between ASPN and the Nrf2 structure. The results revealed that ASPN and Nrf2 tightly bound through 8 hydrogen bonds (Fig. 5C), with docking and confidence scores indicating stable affinity (Figure S8).

Fig. 5.

ASPN may play a role by interacting with Nrf2 transcription factor. (A, B) The DEGs discovered after high-throughput sequencing were visually represented by heat maps and volcano plots, and Nfe2l2 is significantly upregulated after Cap-Exo/ASPN-Cas9 treatment. (C) Molecular docking results of ASPN and Nrf2. (D) Immunoprecipitation showed the interaction between ASPN and Nrf2. (E) WB was used to detect the expression level of total Nrf2 under normal conditions and OA conditions. (F) WB was used to detect the expression levels of nuclear and cytoplasmic Nrf2 in normal state and OA condition with or without ASPN knockout. (G) ARE-luciferase reporter assay to detect the effect of ASPN knockdown on Nrf2 transcription. (H, I) KEGG and GO analysis of DEGs. (J) GSEA enrichment analysis of ECM organization, cartilage development, cell senescence and ferroptosis between Cap-Exo/ASPN-Cas9 treated and control groups

To further confirm this interaction, we used immunoprecipitation technology to prove that ASPN interacts with Nrf2 in vitro, thus speculating that ASPN exerts its effect by binding to Nrf2 (Fig. 5D). The results of WB experiments found no significant difference in the total Nrf2 expression of the two groups of si-NC and si-ASPN under control and TBHP conditions, suggesting that ASPN knockout did not affect the total Nrf2 expression under normal and OA conditions (Fig. 5E). In order to further verify that ASPN may exert its transcriptional function by affecting the entry of Nrf2 into the nucleus, rather than affecting the expression and degradation of Nrf2, we had to conduct a nuclear-cytoplasmic separation experiment. The results, as shown in Fig. 5F, show that under normal and TBHP conditions, after ASPN knockout, the amount of Nrf2 in the nucleus increased and the amount of Nrf2 in the cytoplasm decreased, indicating that ASPN binds to Nrf2 in the cytoplasm, resulting in a decrease in the entry of Nrf2 into the nucleus, further affecting the downstream mechanism. We also conducted additional experiments to directly assess whether ASPN affects Nrf2 transcriptional activity. Specifically, we performed ARE-luciferase reporter gene assays. We examined the effect of ASPN knockdown by comparing the si-NC group and the si-ASPN group (Fig. 5G). The results showed that silencing ASPN significantly increased ARE-luciferase activity, indicating enhanced Nrf2 transcriptional activity.

These observations suggest a strong physiological association between ASPN and Nrf2, where they are typically co-degraded. In contrast, under OA conditions, increased ASPN levels result in accelerated Nrf2 degradation. After determining the crucial role of Nrf2 signaling in mediating ASPN-related effects, we further employed GO and KEGG enrichment analyses to investigate biological processes linked to the identified DEGs. The top 10 GO categories across molecular function, cellular component, and biological process domains were highlighted, suggesting that Cap-Exo/ASPN-Cas9 might play a significant role in inhibiting ferroptosis and maintaining ECM homeostasis (Fig. 5H). KEGG analysis of the top 20 pathways also highlighted that ferroptosis and cellular senescence play key roles in Cap-Exo/ASPN-Cas9 treatment (Fig. 5I). GSEA indicated an upregulation in ECM organization and cartilage development, whereas pathways associated with cellular senescence and ferroptosis were notably downregulated (Fig. 5J). In conclusion, the reduction in Nrf2 degradation following ASPN knockout may further exert its effects by inhibiting ferroptosis and cellular senescence, thereby helping to maintain ECM integrity in OA.

Cap-Exo/ASPN-Cas9 inhibits ferroptosis in chondrocytes and alleviates cellular senescence in vitro

To further clarify how Cap-Exo/ASPN-Cas9 influences ferroptosis and cellular senescence, supplemental experiments were undertaken in vitro. Ferroptosis, an iron-dependent programmed cell death pathway, prominently involves lipid peroxidation [20]. Intracellular ROS, critical ferroptosis markers, were measured using fluorescence-based indicators (DCFH-DA and DHE). Results revealed markedly enhanced ROS generation in chondrocytes after TBHP treatment, while subsequent application of Cap-Exo/ASPN-Cas9 effectively reduced ROS concentrations to levels comparable with ferroptosis inhibitor Ferrostatin-1 (Fer-1), thereby confirming the protective potential of this therapeutic strategy (Fig. 6A, B). Since ferroptosis is intrinsically linked to lipid peroxidation and excessive iron accumulation, we further evaluated ferroptosis markers by measuring lipid peroxidation and iron content in chondrocytes treated with Cap-Exo/ASPN-Cas9. The study demonstrated that TBHP treatment significantly enhanced the fluorescence of the orange iron probe, specific for Fe²⁺ detection, and resulted in a substantial accumulation of lipid ROS, as evidenced by increased green fluorescence. However, compared to conventional exosomes and non-cartilage-targeted exosomes, Cap-Exo/ASPN-Cas9 and Fer-1 treatment significantly reduced the fluorescence intensity of both the iron probe and lipid peroxidation (Fig. 6A, B). Subsequently, we performed WB analysis to evaluate the abundance of proteins related to ferroptosis. In the in vitro OA model, TBHP treatment led to decreased expression of SLC7A11, GPX4, and FTH1, whereas ACSL4 expression increased. Cap-Exo/ASPN-Cas9 and Fer-1 treatment notably counteracted these effects by upregulating SLC7A11, GPX4, and FTH1 expression, thereby preventing TBHP-induced ferroptosis in chondrocytes (Fig. 6C, D). IF imaging revealed that TBHP treatment markedly reduced the expression of ferroptosis-related proteins SLC7A11 and GPX4 in chondrocytes. However, treatment with Cap-Exo/ASPN-Cas9 restored the fluorescence intensity of these proteins, demonstrating a significant protective effect (Fig. 6E, F, S9).

Fig. 6.

ASPN knockout may inhibit ferroptosis and alleviate cell senescence. (A) DHE and DCFH-DA fluorescent probes were used to detect intracellular ROS, the orange iron probe was used to detect intracellular Fe²⁺ levels, and the BODIPY-C11 fluorescent probe was used to visualize lipid peroxidation levels. (B) Quantitative analysis of intracellular ROS, Fe²⁺ levels, and lipid peroxidation levels. (C, D) WB was used to detect the expression of ferroptosis-related proteins. (E, F) Representative IF images of SLC7A11 (red) and GPX4 (green) in chondrocytes after treatment with different groups. (G, H) WB technique was used to detect the expression of Senescence-associated proteins. (I, J) β-galactosidase staining in chondrocytes after treatment in different groups can be used to evaluate the degree of cell senescence. (n = 3, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001)

Cellular senescence refers to a state where cells cease to divide and enter irreversible cell cycle arrest after experiencing stress or damage [21]. Ferroptosis-related DNA damage, cell cycle arrest, and senescence characteristics may expedite aging. We evaluated Cap-Exo/ASPN-Cas9’s impact on cellular senescence by analyzing senescence-associated protein expression using WB. The findings demonstrated that treatment with TBHP markedly enhanced the expression of senescence-related proteins (P53, P21, and P16), whereas Cap-Exo/ASPN-Cas9 administration substantially attenuated this induction (Fig. 6G, H). In agreement, qPCR analysis confirmed significant repression of senescence-associated gene expression following Cap-Exo/ASPN-Cas9 treatment (Figure S10). Additionally, cellular senescence was further assessed by measuring senescence-associated β-galactosidase (SA-β-Gal) activity, a widely recognized biomarker that indicates the progression of cell aging [22]. The findings indicated that TBHP treatment significantly elevated SA-β-Gal activity, indicating a pronounced senescent phenotype, whereas Cap-Exo/ASPN-Cas9 treatment substantially decreased SA-β-Gal expression (Fig. 6I, J). Together, our findings demonstrate that Cap-Exo/ASPN-Cas9 effectively inhibits ferroptosis and lipid peroxidation in chondrocytes, suppresses cellular senescence, and prevents the development of senescent phenotypes in vitro.

Cap-Exo/ASPN-Cas9 alleviates mitochondrial dysfunction by inhibiting ferroptosis through activating the Nrf2/HO-1 pathway

Our previous studies identified Nrf2 as a key mediator of ASPN’s action. When ASPN is knocked out, the degradation of Nrf2 is reduced, and the amount of free Nrf2 in the nucleus increases, which may be crucial for its subsequent effects. The Nrf2/HO-1 pathway is crucial for cellular defense against oxidative stress and environmental challenges, and its association with ferroptosis has garnered growing interest [23]. To investigate if ferroptosis inhibition mediated by Cap-Exo/ASPN-Cas9 involves activation of the Nrf2/HO-1 pathway, we utilized WB assays to analyze related protein expressions. The experimental results demonstrated that nuclear localization of Nrf2 (n-Nrf2) decreased markedly upon TBHP-induced ROS elevation. In contrast, Cap-Exo/ASPN-Cas9 administration effectively recovered n-Nrf2 expression and enhanced downstream antioxidant proteins, including HO-1 and NQO1, thereby improving cellular antioxidative capacity (Fig. 7A, B). These findings were corroborated by IF analysis, which demonstrated pronounced nuclear accumulation of Nrf2 following Cap-Exo/ASPN-Cas9 exposure, reflected by significantly increased nuclear fluorescence intensities (Figure S11). We further performed ARE-luciferase assays on five experimental groups (consistent with our main experimental design), and the results were consistent with the above findings, indicating that the regulation of ASPN expression leads to corresponding changes in Nrf2 transcriptional activity, which also supports the conclusion that Nrf2 accumulates significantly in the cell nucleus (Fig. 7C). Additionally, ML385, a specific Nrf2 inhibitor, was employed to confirm changes in proteins associated with ferroptosis [24]. ML385 partially reversed Cap-Exo/ASPN-Cas9-induced alterations in ferroptosis markers, including GPX4 and SLC7A11 (Figure S12A, B). These observations collectively imply that ASPN downregulation activates the Nrf2/HO-1 signaling axis, thereby inhibiting ferroptotic cell death.

Fig. 7.

ASPN knockout inhibits ferroptosis and mitochondrial dysfunction by activating the Nrf2/HO-1 pathway. (A, B) WB technique was used to detect the expression of Nrf2-related pathway proteins. (C) ARE-luciferase assay to assess whether ASPN affects Nrf2 transcriptional activity. (D)Representative TEM images of mitochondria after treatment in different groups. (E, F) WB was used to detect the expression of proteins related to mitochondrial function. (G) MitoSox probe was used to detect the production of mitochondrial superoxide, MitoTracker Red was used to specifically label biologically active mitochondria, and JC-1 probe was used to sensitively detect changes in mitochondrial membrane potential. (H) Quantitative analysis of MitoSox probe, MitoTracker Red and JC-1 probe. (n = 3, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001)

Mitochondria, as the energy factories and metabolic centers of cells, are also the primary sites for ROS generation and play a pivotal role in ferroptosis. Ferroptosis is characterized by damage to both the outer and inner mitochondrial membranes. TEM analysis validated these findings, revealing notable mitochondrial swelling and compromised membrane integrity following TBHP treatment. However, Cap-Exo/ASPN-Cas9 treatment greatly alleviated these abnormalities (Fig. 7D). Mitochondrial dynamics, encompassing fusion, fission, morphology, and function, are essential for sustaining cellular metabolism, stress response, and cell death regulation. OPA1, DRP1, MFN1, and MFN2 are essential proteins that play core roles in mitochondrial function [25]. WB analysis revealed that Cap-Exo/ASPN-Cas9 treatment elevated OPA1, MFN1, and MFN2 expression levels and reduced DRP1 expression, suggesting its potential in alleviating TBHP-induced mitochondrial damage (Fig. 7E, F).

We also employed MitoSox, a probe that specifically targets mitochondria to detect superoxide generation and dismutation. The results showed that TBHP treatment significantly increased ROS levels in the cells, whereas Cap-Exo/ASPN-Cas9 treatment markedly reduced this effect. MitoTracker staining demonstrated that TBHP impaired mitochondrial activity in chondrocytes, but ASPN-targeted knockout reversed these changes induced by TBHP (Fig. 7G, H). We employed JC-1 staining to evaluate mitochondrial membrane potential. JC-1, an amphipathic fluorescent dye, enters mitochondria and forms red fluorescent polymers when membrane potential is high. In mitochondria with low membrane potential, JC-1 remains as a green-fluorescent monomer. The results showed that TBHP treatment significantly reduced mitochondrial membrane potential, while Cap-Exo/ASPN-Cas9 treatment greatly restored mitochondrial membrane potential to levels close to baseline (Fig. 7G, H). In conclusion, ASPN knockout activates the Nrf2/HO-1 pathway, inhibiting ferroptosis in chondrocytes, mitigating mitochondrial dysfunction, reducing inflammation and ECM degradation, and decreasing cellular senescence.

CRISPR-Cas9-mediated ASPN knockout alleviates OA in mice in vivo

To evaluate the in vivo therapeutic efficacy of CRISPR–Cas9–mediated ASPN deletion and compare it with celecoxib (an FDA-approved OA medication), we established an OA mouse model using DMM surgery in 8-week-old male C57BL/6 mice. Animals then received weekly intra-articular injections of Exo, Cap-Exo, Exo/ASPN-Cas9, Cap-Exo/ASPN-Cas9, celecoxib, or PBS for four weeks, after which joint tissues were collected for analysis (Fig. 8A). Histopathological examination of major organs (heart, lungs, liver, kidneys, spleen) revealed no detectable systemic toxicity associated with Cap-Exo/ASPN-Cas9 delivery (Figure S13). X-ray and microCT assessments showed that DMM-induced joints displayed pronounced osteophyte formation, subchondral bone sclerosis, trabecular abnormalities, and joint space narrowing (Fig. 8B–C). Exo treatment provided mild improvement, whereas Cap-Exo and Exo/ASPN-Cas9 produced stronger but comparable benefits. Notably, Cap-Exo/ASPN-Cas9 achieved the most pronounced protection, markedly reducing osteophytes, smoothing joint surfaces, increasing trabecular density, and expanding joint space—effects approaching those of celecoxib.

Fig. 8.

Cartilage-targeted engineered exosomes alleviate osteoarthritis in mice. (A) Schematic diagram of the in vivo treatment strategy in mice. (B) Representative images of knee joint X-ray and Micro-CT3D reconstruction of mice in different treatment groups. The yellow arrows represent the narrowing of the joint space and the yellow dotted circles represent the formation of osteophytes. (C) Quantitative analysis of osteophyte maturity, osteophyte size, bone volume fraction, and trabecular thickness. (D) Histological staining of mouse knee joint sections, including safranin fast green, TB and H-E staining. (E, F) OARSI score and synovial score of mouse knee joint histological staining. (n = 6, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001)

Moreover, SO and TB staining found that cartilage in the DMM group was significantly worn and the cartilage thickness was reduced, with an OARSI (Osteoarthritis Research Society International) score of 5.67 points. After Exo treatment, cartilage degeneration was slightly alleviated, and OARSI was reduced to a certain extent, about 4.17 points, but it was also weaker than the effects of the Cap-Exo and Exo/ASPN-Cas9 groups. It is worth noting that Cap-Exo/ASPN-Cas9 treatment can significantly antagonize cartilage degradation, promote cartilage thickness recovery, and improve the OARSI score in the DMM model, with a score of up to 1.33 points, while the OARSI score of FDA-approved celecoxib is only 1.00 points. Although there are differences between the two, the differences are very small. The H&E staining results of the synovium revealed that the degree of synovitis was the most obvious in the DMM group, with the highest synovitis score of 5.17 points. Exo, Cap-Exo and Exo/ASPN-Cas9 could inhibit synovitis to a certain extent, with scores of 3.67, 2.83 and 2.83 points, respectively. However, Cap-Exo/ASPN-Cas9 treatment could significantly alleviate knee synovitis, with the score significantly reduced by about 2.00 points (Fig. 8D, E, F).

Additionally, we performed immunohistochemistry to further validate the effects of Cap-Exo/ASPN-Cas9 in the OA mouse model, particularly its role in activating the Nrf2 pathway to inhibit ferroptosis and mitigate mitochondrial dysfunction, thus alleviating inflammation, ECM degradation, and cellular senescence. As depicted in Figs. 9A and S14, the DMM procedure significantly reduced Nrf2-positive regions, substantially increased ACSL4-positive regions (a ferroptosis indicator), elevated MMP-13 expression, and reduced type II collagen levels. Furthermore, increased expression of inflammatory markers, including iNOS and COX2, was observed. Although Exo alone moderately improved these pathological changes, the therapeutic effects remained limited. The use of Cap-Exo and Exo/ASPN-Cas9 has been further improved on the basis of Exo, but it is worth noting that after the introduction of Cap-Exo/ASPN-Cas9, the therapeutic effect further significantly reversed these negative effects on the basis of Cap-Exo and Exo/ASPN-Cas9, and even to some extent was comparable to celecoxib, which is consistent with our in vitro experimental results. IF staining of cartilage tissue revealed a substantial reduction in GPX4 fluorescence intensity (0.23-fold relative to control) in the DMM model, indicative of enhanced ferroptosis. Treatment with Cap-Exo/ASPN-Cas9 markedly restored GPX4 levels to approximately 0.79-fold relative to control. Importantly, this restorative effect was comparable to celecoxib treatment, where GPX4 intensity reached 0.84-fold relative to control, indicating effective suppression of chondrocyte ferroptosis (Fig. 9B, C). The anti-inflammatory effect of Cap-Exo/ASPN-Cas9 in vivo is shown in Figure S15A, which can greatly inhibit the expression of CD68 in the synovium, which is a driving factor for arthritic joint destruction. In addition, Cap-Exo/ASPN-Cas9 can also reduce the expression of DRP1, which is one of the most important proteins that determine mitochondrial fission and determines the balance of mitochondrial dynamics. The fluorescence results confired DRP1 in the DMM group was increased, and the treatment of Cap-Exo/ASPN-Cas9 can greatly reduce the expression of DRP1 compared with Exo and Exo/ASPN-Cas9, which also indicates that it alleviates mitochondrial dysfunction in vivo (Figure S15B, C). Moreover, the expression of senescence marker P16, often referred to as the “senescence switch”, was significantly elevated (2.19-fold) in the DMM group compared with controls, confirming that cellular senescence pathways are activated during OA development. Notably, Cap-Exo/ASPN-Cas9 intervention significantly reduced P16 expression levels, thereby alleviating chondrocyte senescence (Fig. 9D, E).

Fig. 9.

Cap-Exo/ASPN-Cas9 can activate Nrf2 in mice to inhibit ferroptosis and mitochondrial dysfunction and alleviate OA. (A) Immunohistochemistry was used to detect the expression of Nrf2, ACSL4, MMP-13 and Collagen-II in cartilage, and COX2 and INOS in synovial tissue. (B, C) Representative IF images and quantitative analysis of GPX4 in the knee cartilage of mice after treatment with different groups. (D, E) Representative IF images and quantitative analysis of P16 in the knee cartilage of mice after treatment with different groups. (n = 6, independent samples, Mean ± SD, one-way ANOVA, Tukey’s multiple comparisons test, ns, no significant difference; *p < 0.05; **p < 0.01; ***p < 0.001)

Discussion

Osteoarthritis (OA) is a progressive joint disorder marked by synovial inflammation, cartilage degradation, and subchondral bone changes, affecting millions worldwide. Its pathogenesis involves the interaction of multiple factors, including biomechanics, inflammation, and metabolism. Increasing evidence suggests that oxidative stress-induced chondrocyte dysfunction, mitochondrial damage, ferroptosis, and cellular senescence are not independent events, but rather constitute an interconnected pathological network that accelerates the progression of osteoarthritis (OA) [26, 27]. However, the upstream molecular regulators coordinating these processes remain not fully elucidated. Current therapies—ranging from analgesics to joint replacement—mainly provide symptomatic relief and do not target the molecular drivers of disease progression [28].

Asporin (ASPN), a leucine-rich repeat-containing extracellular matrix protein highly expressed in cartilage, has emerged as a critical mediator of OA pathogenesis. Elevated ASPN expression correlates with disease severity and contributes to extracellular matrix degradation and inflammatory amplification [29, 30]. In the present study, we identify ASPN as a key upstream regulator linking redox imbalance, ferroptosis, mitochondrial dysfunction, and chondrocyte senescence. By employing Cap-Exo/ASPN-Cas9–mediated CRISPR-Cas9 knockout of ASPN, we demonstrate that targeting ASPN can simultaneously modulate multiple pathological processes central to OA progression.

Mechanistically, our findings highlight nuclear factor erythroid 2–related factor 2 (Nrf2) as a pivotal downstream effector of ASPN. Nrf2 is a master transcriptional regulator of cellular antioxidant defense and redox homeostasis, controlling the expression of genes such as HO-1, GPX4, and other cytoprotective enzymes [31]. In osteoarthritis, reduced Nrf2 nuclear translocation and excessive reactive oxygen species (ROS) exceed antioxidant capacity, leading to oxidative damage and cartilage degeneration [32, 33]. Our data indicate that ASPN directly interacts with Nrf2 in the cytoplasm, limiting its nuclear translocation and transcriptional activity. This interaction suppresses Nrf2-mediated antioxidant signaling, thereby rendering chondrocytes more vulnerable to oxidative stress. Importantly, ASPN knockout markedly reduced ASPN–Nrf2 binding, restored Nrf2 nuclear accumulation, and enhanced antioxidant responses, positioning ASPN as an upstream negative regulator of Nrf2 signaling in OA.

Suppression of Nrf2 activity has profound downstream consequences, particularly in the regulation of ferroptosis. Ferroptosis is an iron-dependent form of regulated cell death characterized by lipid peroxidation and depletion of antioxidant defenses, and is increasingly recognized as a major contributor to OA pathogenesis [34]. Excess iron accumulation and reduced expression of key ferroptosis suppressors such as GPX4 and GSH have been reported in OA cartilage and synovial fluid [35]. In line with these observations, we found that ASPN knockout via Cap-Exo/ASPN-Cas9 significantly attenuated lipid peroxidation and iron overload, thereby inhibiting ferroptosis through reactivation of the Nrf2/HO-1 antioxidant axis. These findings establish a mechanistic link between ASPN-mediated Nrf2 suppression and ferroptotic vulnerability in chondrocytes.

Mitochondrial dysfunction represents a critical convergence point between oxidative stress and ferroptosis. Mitochondria are both a major source and a primary target of ROS, and their impairment exacerbates redox imbalance, lipid peroxidation, and energy failure in OA chondrocytes [36, 37]. Ferroptosis is accompanied by characteristic mitochondrial abnormalities, including mitochondrial shrinkage, increased membrane density, and collapse of membrane potential [38]. In this study, Cap-Exo/ASPN-Cas9 treatment substantially restored mitochondrial integrity, as evidenced by normalized mitochondrial morphology, improved membrane potential, and reduced ROS accumulation. These effects were accompanied by rebalancing of mitochondrial dynamics, with increased expression of fusion-related proteins (OPA1, MFN1, MFN2) and suppression of the fission regulator DRP1 [39]. By preserving mitochondrial homeostasis, ASPN knockout disrupts a key source of lipid ROS generation, thereby further restraining ferroptotic signaling.

Persistent oxidative stress, ferroptosis, and mitochondrial dysfunction collectively drive chondrocyte senescence, a hallmark of OA progression [40]. Senescent chondrocytes undergo irreversible cell cycle arrest and secrete pro-inflammatory cytokines and matrix-degrading enzymes, collectively termed the senescence-associated secretory phenotype (SASP), which accelerates cartilage degeneration and synovial inflammation [41, 42]. Emerging evidence indicates that excessive ROS production, mitochondrial damage, and lipid peroxidation activate senescence-associated pathways such as p16 and p21, forming a self-amplifying loop between cellular aging and tissue degeneration [43]. Consistent with this model, Cap-Exo/ASPN-Cas9 treatment significantly reduced SA-β-Gal–positive cells and downregulated p16 and p21 expression. These anti-senescent effects coincided with reduced ferroptotic stress and restored mitochondrial function, suggesting that disruption of the ASPN–Nrf2–ferroptosis axis effectively alleviates senescence-driven cartilage deterioration.

Collectively, our findings support a unified pathogenic model in which ASPN acts as an upstream molecular switch that suppresses Nrf2 signaling, thereby promoting ferroptosis, mitochondrial dysfunction, and chondrocyte senescence in OA. By targeting ASPN using an exosome-based CRISPR-Cas9 delivery platform, this pathological cascade can be simultaneously interrupted, offering a multifaceted therapeutic strategy to preserve cartilage homeostasis and slow OA progression.

However, several limitations of this study warrant further investigation. First, although Cap-Exo/ASPN-Cas9 significantly reduced ASPN expression, the loading efficiency and stability of the exosomal delivery strategy still require further optimization. Second, despite the improved delivery specificity conferred by exosomes, challenges such as limited cartilage penetration, synovial clearance, and insufficient retention within deep cartilage zones remain unresolved. Future engineering strategies that enable simultaneous tissue- and cell-specific targeting—such as surface charge modulation, incorporation of matrix-penetrating peptides, or hydrogel-based delivery systems—may further enhance intra-articular retention and therapeutic efficacy. Importantly, while our data indicate that ASPN interacts with Nrf2 and regulates its transcriptional activity, the exact molecular mechanisms of this interaction remain incompletely elucidated. Although we designed Nrf2 truncated mutants, this study did not complete comprehensive binding domain localization experiments. Such domain-level resolution will provide crucial mechanistic insights into the regulation of redox homeostasis and stress responses during osteoarthritis progression and may contribute to the development of domain-specific inhibitors or decoy strategies to selectively restore Nrf2 activity. Therefore, identifying the specific Nrf2 domains responsible for ASPN binding is crucial for elucidating how this interaction interferes with Nrf2 transcriptional activity and downstream antioxidant signaling pathways in chondrocytes, and further validation is needed in future mechanistic studies. In addition, although both in vitro and in vivo assays confirmed the short-term safety and biocompatibility of Cap-Exo/ASPN-Cas9, more comprehensive evaluations are required to assess its immunogenicity, potential macrophage activation, and long-term joint toxicity. Studies in large animal models, together with the development of scalable and standardized production processes, will be essential to facilitate future clinical translation.

Conclusion

In conclusion, this study demonstrates that Cap-Exo/ASPN-Cas9 provides significant protection against OA both in vivo and in vitro. ASPN knockout effectively reduces Nrf2 degradation, activates the Nrf2/HO-1 pathway, and inhibits ferroptosis in chondrocytes. Additionally, Cap-Exo/ASPN-Cas9 treatment alleviates mitochondrial dysfunction, diminishes inflammation, maintains chondrocyte microenvironmental homeostasis, reduces cellular senescence, and retards OA progression. Thus, targeted CRISPR-Cas9-mediated deletion of ASPN represents a novel and promising approach for OA therapy, establishing a foundation for future treatment strategies.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Chao Lou: Writing-original draft. Jinwu Wang: Methodology, Formal analysis. Chengqian Dai: Investigation, Formal analysis. Jilong Wang and Jin Yang: Investigation. Yuqin Fang: Software. Hongyi Jiang: Conceptualization. Xiaoyun Pan: Investigation. Han Li and Chenhao Lan: Methodology, Investigation. Guohong Xu: Validation. Shoaib Iqbal: Supervision, Validation. Jiaqian Bao: Supervision. Leyi Cai: Supervision, Methodology, Project administration. Wenhao Zheng: Conceptualization, Funding acquisition.

Funding

This work was funded by the Basic Public Welfare Research Project of Zhejiang Province (LQ24H060008), Zhejiang Medicine Health Science and Technology Program (2025HY0585) and Wenzhou Science and Technology Plan Project (GY20250280).

Data availability

The data are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

The Animal Ethics Committee of Wenzhou Medical University approved this study (wydw2025-0032).

Consent for publication

All authors agreed to the publication of this manuscript.

Competing interests

The authors declare no competing interests.