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Journal of Extracellular Vesicles logoLink to Journal of Extracellular Vesicles
. 2026 Apr 7;15(4):e70261. doi: 10.1002/jev2.70261

Cisplatin Promotes Immunosuppression in Ovarian Cancer by Enhancing miR‐181a‐5p‐Enriched Extracellular Vesicles to Drive Regulatory T Cell Differentiation

Liyan Zheng 1,2, Sijia Mu 1,3, Mingxue Liu 4, Panmin Lei 1, Jingchen Zhang 1, Yuting Dai 1, Chenxi He 1,3, Fengli Qu 1,4,✉, Weihong Tan 1,2
PMCID: PMC13054833  PMID: 41944372

ABSTRACT

Although cisplatin (CDDP) is widely employed in combination immunotherapy, no CDDP‐based regimen has shown a survival benefit when treating the ovarian cancers. This is mainly because the impact of CDDP on immunotherapy is not fully understood. A critical gap concerns how CDDP reshape the tumour microenvironment, especially through extracellular vesicles (EVs)‐mediated communication. In this work, using human and murine ovarian cancer cells as a model, we demonstrate that CDDP boosts the secretion of EVs from cancer cells, while exerting no such effect on non‐cancerous cells. These CDDP‐induced tumour‐derived EVs, in turn, drive the differentiation of CD4+ T cells towards immunosuppressive regulatory T cells (Treg cells), which are known to limit the efficacy of immunotherapy. Based on next‐generation sequencing, a significant enrichment of miR‐181a‐5p was identified in CDDP‐induced tumour‐derived EVs, and further functional studies confirmed that this microRNA promoted Treg cell differentiation via suppressing sirtuin 1 (SIRT1), a key regulator of the transcription factor forkhead box protein P3 (FOXP3). Importantly, inhibition of miR‐181a‐5p abrogated the Treg‐promoting effect of CDDP‐induced tumour‐derived EVs, a finding further validated in vivo, where blockade of miR‐181a‐5p not only impaired Treg differentiation but also restored T‐cell‐mediated antitumour immunity and restrained tumour growth. Together, these findings uncover a previously unrecognised mechanism by which CDDP exacerbates immunosuppression via miR‐181a‐5p‐enriched EVs and suggest that targeting this pathway could improve the therapeutic efficacy of combination immunotherapy in ovarian cancer.

Keywords: cisplatin, extracellular vesicles, microRNAs, ovarian cancer

Short abstract

Herein, we find that CDDP induces ovarian cancer cells to secrete miR‐181a‐5p‐enriched EVs, which subsequently enhances the differentiation of CD4+ T cells towards Treg cells via SIRT1‐FOXP3 signalling pathway. By blocking miR‐181a‐5p‐enriched tumour‐derived EVs, the differentiation of Treg cells is abolished, thus offering a potential target for enhancing immunotherapy in ovarian cancer.

1. Introduction

Cisplatin (CDDP), administered either as monotherapy or in neoadjuvant regimens, remains the most widely used chemotherapeutic agent for ovarian cancer, which exhibits the highest mortality among gynaecological malignancies owing to its frequent relapse and metastasis (McGuire and Markman 2003; Vergote et al. 2010; Lawrie et al. 2015). In recent years, novel CDDP/immune checkpoint inhibitor (ICI) has been evaluated in treating ovarian cancer, aiming to exploit synergism by host immune response for improved overall survival (Konstantinopoulos and Cannistra 2021; Lo et al. 2017). However, first‐line clinical trials have not shown a survival advantage, possibly because the effects of CDDP on tumour immune microenvironment have not been fully addressed (Monk et al. 2021; Moore et al. 2021; Curiel et al. 2004). Accumulated evidence in various cancers suggests that many chemotherapeutic agents can promote the differentiation and infiltration of intratumoural immunosuppressive regulatory T cells (Treg cells), either directly or indirectly, thus hindering T cell effector immune response (Lee et al. 2021; Nishikawa and Sakaguchi 2014). Despite this knowledge, the underlying mechanism by which CDDP affects Treg cells in the context of optimising combination regimens for ovarian cancer remains to be elucidated.

Extracellular vesicles (EVs), a subset of cell‐derived biological vesicles, play a crucial role in intercellular communication by transporting signalling molecules, such as nucleic acids, proteins and lipids, from parent cells to adjacent or distant recipient cells (Kalluri and LeBleu 2020). Numerous reports have pointed out that EVs can modulate the behaviour of recipient cells, with tumour‐derived EVs (TEVs) receiving particular attention due to their involvement in tumour development, metastasis and relapse (Keklikoglou et al. 2018; Fang et al. 2018; Zhang and Yu 2019; He et al. 2015). For instance, TEVs have been implicated in regulating pre‐metastatic niche microenvironment and promoting lung metastasis of highly metastatic hepatocellular carcinoma through interacting with fibroblasts (Keklikoglou et al. 2018). In addition, TEVs have been associated with poor immune response in diverse malignancies, as they can inhibit host antitumour responses and drive drug resistance by modulating the differentiation and infiltration of intratumoural Treg cells (Wu et al. 2023). According to the clinic data, the amount of TEVs was found to be positively correlated with that of intratumoural CD4+ Treg cells and inversely correlated with the efficacy of immunotherapy (Wei et al. 2023; Xu et al. 2020). Given the limited success of CDDP/ICI combination in clinical trials for ovarian cancer, we hypothesise that CDDP might induce pro‐cancer TEVs that are involved in the expansion of immunosuppressive Treg cells, thereby leading to immune suppression.

In this work, we investigate the mechanisms underlying the limited efficacy of combining CDDP with ICI in first‐line clinical trials for ovarian cancer. Using human (SKOV3) and murine (ID8) ovarian adenocarcinoma cell lines, together with a non‐cancerous ovarian epithelial cell line (IOSE80), we found that CDDP significantly enhanced EV secretion from ovarian cancer cells but not from non‐cancerous cells. These CDDP‐induced TEVs further promoted the differentiation of CD4+ T cells into Treg cells. Furthermore, next‐generation sequencing revealed a significant enrichment of miR‐181a‐5p in CDDP‐induced TEVs, which was validated as a key mediator of Treg differentiation by activating forkhead box protein P3 (FOXP3) expression via suppressing sirtuin 1 (SIRT1, a well‐known negative regulator of Treg cells). Both in vitro and in vivo studies confirmed that inhibition of miR‐181a‐5p reduced Treg differentiation, restored cytotoxic T cell function and enhanced antitumour efficacy. Notably, blockade of miR‐181a‐5p‐enriched EVs effectively abolished CDDP‐enhanced FOXP3 upregulation and Treg cell expansion. Collectively, our findings reveal a previously unrecognised mechanism by which CDDP exacerbates immunosuppression through miR‐181a‐5p‐enriched TEVs and identify this pathway as a promising therapeutic target to improve the efficacy of combination immunotherapy in ovarian cancer.

2. Materials and Methods

2.1. Materials and Reagent s

All oligonucleotide sequences were purchased from Sangon Biotech, Shanghai. The cell counting kit‐8 (CCK‐8) was purchased from Dojindo, Japan. Phenylmethanesulfonyl fluoride (PMSF) solution and 3,3′‐dioctadecyloxacarbocyanine perchlorate (DiO) were purchased from Beyotime, China. RIPA lysis buffer was obtained from Applygen, China. Antibodies used for flow cytometry, including PE anti‐human FOXP3 antibody, APC anti‐human CD25 antibody, PE/Cyanine7 anti‐human CD4 antibody, FITC anti‐human CD3 antibody, Brilliant Violet 421 anti‐human CD8 antibody, PE anti‐mouse/rat/human FOXP3 antibody, FITC anti‐mouse CD25 antibody, FITC anti‐mouse CD3 antibody, APC anti‐mouse CD4 antibody and PE anti‐mouse CD8a antibody, were purchased from BioLegend, USA. The Intracellular Fixation & Permeabilization Buffer Set was also obtained from BioLegend. Rabbit monoclonal antibodies against CD63, TSG101, Alix and GM130 (reactive to human or mouse proteins) for western blotting (WB) were purchased from Proteintech, USA. MACS column, MACS Separator, T Cell TransAct, human CD4 MicroBeads and human CD8 MicroBeads were purchased from Miltenyi BioTechnologies Co., LTD. Collagenase IV was purchased from Gibco, USA.

2.2. Cell Lines and Culture Methods

The human ovarian cancer cell line SKOV3 and human non‐cancerous ovarian epithelial cell line IOSE80 were originally obtained from American Type Culture Collection. The mouse ovarian cancer cell line ID8‐luciferase (ID8‐LUC) was originally obtained from MeisenCTCC. All cells were maintained in DMEM medium supplemented with 10% fetal bovine serum (FBS), 1% penicillin–streptomycin (PS) and cultured at 37°C in a humidified incubator (Thermo Fisher Scientific) with 5% CO2 and 95% humidity. CD4+ and CD8+ T cells were isolated from healthy donor peripheral blood mononuclear cells and cultured according to the protocol of commercial kits. The use of human samples was approved by the relevant Institutional Review Board (approval number: IRB‐2024‐87 (IIT)).

2.3. Cell Viability

Cell viability was assessed using a CCK‐8 assay. Briefly, attached cells were seeded in a 96‐well plate and incubated overnight in 100 µL of DMEM supplemented with 10% FBS and 1% PS. Then, the drugCDDP was added at increasing concentrations and incubated for 24 h. After this incubation period, 10% (v/v) of the CCK‐8 reagent was added to each well. After 4 h of incubation, the absorbance was measured at 450 nm.

2.4. EVs Isolation

EVs from all cell lines were isolated using the following procedure. In detail, cells were cultured in fresh DMEM supplemented with 10% FBS and 1% PS until they reached approximately 90% confluence. The culture medium was then replaced with serum‐free (FBS‐free) medium, and the cells were incubated for another 48 h. The conditioned medium was collected and subjected to sequential centrifugation: first at 300 × g for 10 min to remove floating cells, followed by 2000 × g for 10 min to pellet dead cells, cellular debris and giant vesicles. The resulting supernatant was then filtered through a 0.22 µm filter and concentrated to 500 µL using a 100‐kDa ultrafiltration tube. Finally, EVs were isolated by size exclusion chromatography for future study. Specifically, after loading 500 µL of the concentrate, the eluate was continuously collected in 500 µL fractions. The third fraction (corresponding to 1.0–1.5 mL of eluate), which was identified as the most EV‐enriched component, was collected for all subsequent experiments. To prepare CDDP‐EVs, cells were stimulated with CDDP for 24 h before being switched to FBS‐free medium to collect EVs following the protocol described above. Purified EVs were quantified using two complementary methods. The particle concentration was analysed by Nanoparticle Tracking Analysis (NTA) using a ZetaView system (Particle Metrix, Germany). The total protein concentration of EVs was determined using a Bicinchoninic Acid (BCA) Protein Assay Kit (Beyotime Biotechnology, China). For subsequent experiments, particle count was used to dose EVs in functional cell assays, while protein amount was used to normalise EV loading for WB analysis.

2.5. Transmission Electron Microscopy (TEM)

A 10‐µL aliquot of the EV suspension (1 × 107 particles/mL) was deposited onto formvar‐carbon‐coated electron microscopy grids for 2 min. The grids were negatively stained with 3% uranyl acetate for 10 s, followed by a second staining step lasting for 1 min. Subsequently, the samples were imaged using a JEM‐2100 Plus Transmission Electron Microscope (JEOL USA, Inc., Peabody, MA) operating at an accelerating voltage of 80 kV.

2.6. Western Blotting

Cells or EVs were lysed on ice for 30 min using RIPA lysis buffer supplemented with 1% PMSF, followed by centrifugation of the lysate at 10,000 × g for 30 min to isolate the proteins. The protein concentration was determined using a BCA assay kit. Subsequently, the protein samples were mixed with 5× protein loading buffer at a 4:1 ratio and denatured at 95°C for 5 min. Then, protein samples (10 µg per lane) were separated on a 4%–20% sodium dodecyl sulfate polyacrylamide gel and subsequently transferred electrophoretically to an immobilon polyvinylidene fluoride membrane (EMD Millipore, Billerica, MA, USA). After incubation with primary antibody overnight at 4°C and subsequent incubation with appropriate horseradish peroxidase‐conjugated secondary antibodies at room temperature for 2 h, the protein bands were visualised using an Amersham ImageQuant 800 imager.

2.7. Nanoparticle Tracking Analysis

EVs were characterised by NTA with a detection range of 0.01–2 µm. Samples were diluted in DPBS to an optimal concentration, and then 1 mL of the dilution was injected into the measurement cell for automatic analysis across 10 positions. Particle size distribution and concentration were determined using the integrated ZetaView software.

2.8. Cellular Uptake

After plating CD4+ T cells in a 4‐well confocal dish at a density of 1 × 105 cells per well, the cells were treated for 24 h with 30 µL of DiO‐labelled EV samples, including EVs and CDDP‐EVs, at 1010 particles/mL. Before analysis, nuclei were stained with Hoechst 33342 for 30 min at 37°C. EV internalisation was finally examined by both confocal laser scanning microscopy (CLSM) and flow cytometry.

2.9. Flow Cytometry Staining

Cell pellets were collected by centrifugation at 300 × g for 4 min. After removing the supernatant, the cells were resuspended in 100 µL of 0.5% BSA solution containing fluorescent antibodies to form a single‐cell suspension, followed by incubation on ice for 20 min in the dark. Subsequently, unbound antibodies were removed by washing the cells twice with DPBS, followed by flow cytometric analysis on an LSR Fortessa instrument (BD Biosciences).

2.10. Transfection With microRNAs (miRNAs) or Anti‐miRNA Mimics

T cells were suspended in FBS‐free medium and seeded in a 24‐well plate with 2 × 106 cells per well, followed by addition of miRNA mimics/anti‐miRNA‐lipofectamine 3000 transfection reagent complexes. The resultant mixture was placed in the incubator at 37°C with 5% CO2 and 95% humidity for 6 h. Next, the transfected T cells of each group were collected by centrifugation, and the cell pellets were resuspended in fresh complete medium and placed in the incubator for further culture.

2.11. Reverse Transcription Quantitative Polymerase Chain Reaction (RT‐qPCR) Analysis

miRNAs were extracted from prepared cells using the EasyPure miRNA Kit (TransGen Biotech, Beijing, China). According to the manufacturer's instructions, the extracted miRNAs were reverse‐transcribed into cDNA using the miRNA First Strand cDNA Synthesis Kit (Sangon Biotech, Shanghai, China). Quantitative polymerase chain reaction (qPCR) was then performed using the 2 × Universal SYBR Green qPCR Master Mix (Sangon Biotech) on a CFX96 Touch Real‐Time PCR Detection System (Bio‐Rad Laboratories, Inc.). The relative expression levels of target miRNAs were normalised to U6 snRNA and calculated using the comparative 2−ΔΔCt method.

2.12. RNase/Protease and Detergent Protection Assay

EVs were divided into four treatment groups: (1) EV group: The intact EV group underwent direct miRNA extraction. (2) EV + RNase A group: EVs were incubated with 100 µg/mL RNase A at 37°C for 1 h, followed by washing via ultracentrifugation and subsequent miRNA extraction. (3) EV + RNase A + Proteinase K group: EVs were first digested with Proteinase K (200 µg/mL, 37°C, 30 min). The protease activity was then inhibited with PMSF, followed by a subsequent treatment with RNase A (100 µg/mL, 37°C, 1 h). Finally, the EVs were washed and subjected to miRNA isolation. (4) EV + 0.1% Triton X‐100 + RNase A group: EVs were prepared by membrane disruption with 0.1% Triton X‐100 on ice for 30 min. The samples were then treated with 100 µg/mL RNase A at 37°C for 1 h. Following digestion, EVs were washed via ultracentrifugation and subjected to miRNA extraction. miR‐181a‐5p levels were quantified by qPCR, with normalisation to the directly extracted intact EV control group.

2.13. Small RNA Sequencing

EVs were isolated from control and CDDP‐treated groups (as described in Section 2.4), with each sample derived from ten 15‐cm culture dishes. The purified EV preparations were submitted to BGI Genomics (Shenzhen, China) and sequenced on the DNBSEQ platform (single‐end 50 bp). Bioinformatic analysis included standard data quality control, alignment to the human reference genome (GRCh38) and quantification of small RNAs.

2.14. In Vivo Therapeutic Efficacy Study

Six‐week‐old female C57BL/6 mice were randomly divided into three groups (n = 3) based on body weight: (i) normal saline (NS), (ii) 1.5 mg/kg CDDP and (iii) 1.5 mg/kg CDDP combined with anti‐miR‐181a‐5p‐loaded EVs (EVs/anti‐miR‐181a‐5p). Intraperitoneal administration was started on Day 18 after the intraperitoneal inoculation with 5 × 106 ID8‐LUC cells. The day of inoculation was recorded as Day 0. The mice were treated every 3 days, which included intraperitoneal injection, body weight measurement and abdominal circumference assessment. Ascites fluid was collected from mice on Days 0, 9 and 24 for subsequent isolation of EVs and cellular analysis. On Day 24, the mice were intraperitoneally injected with 150 mg/kg D‐luciferin and subjected to IVIS imaging. After imaging, the mice were euthanised for terminal analysis. All visible tumour nodules larger than 1 mm in diameter were carefully dissected from the peritoneal cavity, photographed and counted. All animal experiments were conducted in accordance with the guidelines approved by the Animal Ethics Committee of Hangzhou Institute of Medicine, Chinese Academy of Sciences (Approval Number: AP2025‐02‐0458).

2.15. Tumour Immune Microenvironment Analysis

Fresh ascites, tumour tissues and spleens were collected for the detection of relevant immune cells by flow cytometry. Briefly, tumour tissues were minced into small fragments (0.5–1 mm3) and digested in DMEM basal medium containing 2 mg/mL collagenase IV at 37°C for 3 h with shaking. Spleens were mechanically dissociated into single‐cell suspensions using the plunger end of a syringe, followed by erythrocyte lysis buffer treatment for 10 min. Ascites fluid was centrifuged at 3000 × g for 10 min at 4°C. The supernatant was used for EV extraction, while the pellet was treated with erythrocyte lysis buffer for 10 min to collect residual cells. Finally, all cells were filtered through a 300‐mesh nylon sieve to obtain single‐cell suspensions. For tumour and spleen cells, T cells were stained with anti‐mouse FITC‐CD3, APC‐CD4 and PE‐CD8a antibody. Treg cells in ascites were stained with anti‐mouse PE‐FOXP3 and FITC‐CD25 antibody.

2.16. Statistical Analysis

Data are presented as mean ± standard deviation (SD). Group comparisons were performed using t tests in GraphPad Prism 10, with statistical significance defined as p < 0.05. The following symbols indicate levels of significance: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001; ns = not significant.

3. Results

3.1. CDDP Increases EV Secretion From Ovarian Cancer Cells

It has been reported that external factors including chemotherapeutic agents altered the secretion of EVs (Aubertin et al. 2016). To determine whether CDDP impacts the secretion of EVs from cells, ovarian cancer cells (human: SKOV3; murine: ID8) and non‐cancerous ovarian epithelial cells (IOSE80) were exposed to increasing concentrations of CDDP—a chemotherapeutic agent commonly used in standard treatment regimens for ovarian cancer (Chaluvally‐Raghavan et al. 2016; Armstrong et al. 2006). In clinical practice, low‐dose CDDP is frequently administered in combination with other chemotherapeutic agents or alongside radiation therapy to improve treatment efficacy while minimising systemic toxicity. Moreover, the actual drug concentration that reaches and penetrates the tumour site is often low (Kelland 2007). Therefore, a deeper understanding of the interactions between low‐dose CDDP and tumour cells is crucial, as it may lead to optimised dosing strategies that enhance its anticancer activity. To investigate these mechanisms, we first utilised CCK‐8 assays to establish a subtoxic concentration of CDDP that would avoid the confounding influence of significant cell death. This screening identified 0.25 µg/mL as the optimal dose, which was subsequently applied for 24 h in vitro to stimulate EV secretion and function. As shown in Figure S1, such treatment protocol was confirmed to maintain cell viability at 80%–90% without significant growth inhibition in SKOV3, ID8 and IOSE80 cell lines. This approach enables focused analysis of phenotypic adaptations and signalling pathways mediated by ovarian cancer cell‐derived EVs under sustained chemotherapeutic stress.

EVs from the three cell lines, both with and without 24 h CDDP stimulation, were isolated and characterised. As shown in Figure 1a, TEM imaging revealed that both EVs and CDDP‐EVs exhibited the typical cup‐shaped morphology, with no significant difference observed between them. EV concentration measured by NTA showed a comparable particle size distribution between both groups, but a higher particle yielded upon CDDP stimulation (Figure 1b). As shown in Figure 1c–e, WB analysis confirmed the presence of three positive EV markers (ALIX, TSG101 and CD63) in all isolated vesicles, while the negative markers GM130 (a Golgi‐specific protein) and Calnexin (an endoplasmic reticulum protein) were absent. Furthermore, to rule out potential contamination from lipoproteins, we analysed the isolated EVs by WB for the lipoprotein marker ApoA1 and ApoB. As shown in Figure S2, neither ApoA1 nor ApoB was detected in our EV preparations, confirming the absence of significant lipoprotein contamination. Collectively, these results demonstrated the successful isolation of high‐purity EVs and indicated that CDDP treatment had no significant effect on EV morphology, size or the expression of characteristic marker proteins. To ensure the absence of endotoxin contamination in EV preparations, which could otherwise confound immune cell assays, we quantified endotoxin levels using a kinetic turbidimetric Limulus amebocyte lysate assay. A standard curve generated from reference endotoxin (0.1, 1 and 10 EU/mL) showed excellent linearity (y = –0.7075x + 2.8689, R 2 = 0.961). The measured endotoxin concentration in the final EV samples was 0.138 EU/mL (Figure S3), which is substantially below the widely accepted stringent limit for EV studies (<0.25 EU/mL). This result confirms that our EV preparations contain negligible endotoxin and are suitable for subsequent functional experiments with immune cells.

FIGURE 1.

FIGURE 1

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CDDP stimulates EV secretion in ovarian cancer cells. (a) Representative TEM images of EVs and CDDP‐EVs derived from SKOV3, ID8 and IOSE80 cells. Scale bars, 200 nm. (b) Size distribution profiles of EVs and CDDP‐EVs from SKOV3, ID8 and IOSE80 cells, as determined by NTA. (c–e) WB analysis of EV markers (ALIX, TSG101 and CD63) and negative‐control organelle markers (GM130, Calnexin) in lysates of SKOV3 (c), ID8 (d) and IOSE80 (e) cells and their corresponding extracellular vesicle preparations (EVs and CDDP‐EVs). (f) Statistical analysis of EVs released by SKOV3, ID8 and IOSE80 cells treated with or without CDDP, n = 3. CDDP, cisplatin; EV, extracellular vesicle; TEM, transmission electron microscopy.

Next, statistical analysis was applied to the EV concentration data to compare secretion levels among the different cell types. As shown in Figure 1f, the quantity of SKOV3‐CDDP‐EVs and ID8‐CDDP‐EVs was significantly increased compared to that of EVs from untreated groups. In contrast, the amount of CDDP‐EVs from treated IOSE80 cells remained comparable to that of EVs from untreated controls. These findings indicate that CDDP enhances EV secretion in ovarian cancer cells, while exhibiting minimal effect on non‐cancerous ovarian epithelial cells—a phenomenon previously reported in other cancer types but demonstrated here for the first time in ovarian cancer (Bandari et al. 2018).

3.2. CDDP‐Induced TEVs Promote Treg Cell Differentiation

Accumulated evidence demonstrates that TEVs play a critical role in modulating the function of immune cells, especially T cells, to suppress antitumour immune response (Kalluri and LeBleu 2020). Therefore, using SKOV3 cells as a model, we tested whether these CDDP‐induced TEVs (denoted as CDDP‐EVs) influenced the biological functions of CD4+ T and CD8+ T cells, which represented the two most effective subtypes of immune cells for cancer immunotherapy (Antony et al. 2005; Coleman et al. 2005). First, the successful isolation of CD4+ and CD8+ T cells was confirmed before functional assays (Figure S4). Subsequently, all EVs were pre‐stained with DiO and co‐cultured with CD4+ T cells to assess cellular uptake. As shown in Figure 2a, CLSM images confirmed the successful internalisation of both EVs and CDDP‐EVs into CD4+ T cells. Quantitative analysis of the mean fluorescence intensity from 10 randomly selected areas in the CLSM images showed comparable levels between the two groups (Figure 2b). Consistent with this, flow cytometry data (Figure 2c and Figure S5) further demonstrated comparable uptake levels of both EV types by CD4+ and CD8+ T cells, suggesting that CDDP treatment does not alter the internalisation efficiency of EVs. Furthermore, flow cytometric analysis using Annexin V and propidium iodide (PI) staining showed that the apoptotic rates in CD4+ T cells were comparable across all groups, with values of (5.4 ± 0.2)% in the PBS control, (5.4 ± 0.9)% in the EVs group and (4.6 ± 0.1)% in the CDDP‐EVs group (Figure 2d, e). A similar trend was also observed in CD8+ T cells (Figure S6). Collectively, neither EVs nor CDDP‐EVs exerted a significant pro‐apoptotic effect on either CD4+ or CD8+ T cells. This result rules out the possibility that the observed effects of CDDP‐EVs on T cell function were indirectly caused by reduced cell viability.

FIGURE 2.

FIGURE 2

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CDDP‐EVs promote the expansion of Treg cells in vitro. (a) Representative CLSM images showing CD4+ T cells treated with DIO‐stained EVs or DIO‐stained CDDP‐EVs for 24 h. Scale bar, 10 µm. (b) Quantification of the mean DIO fluorescence intensity for each group corresponding to panel a, n = 10. (c) Flow cytometric analysis of CD4+ T cells treated with DIO‐stained EVs or DIO‐stained CDDP‐EVs for 24 h. (d, e) Representative flow cytometry plots (d) and quantification (e) of the apoptotic rates of CD4+ T cells treated with EVs or CDDP‐EVs for 24 h by flow cytometry, n = 3. (f, g) Representative flow cytometry plots (f) and quantification (g) of the CD25+FOXP3+ Treg cells among CD4+ T cells treated with EVs or CDDP‐EVs for 24 h by flow cytometry, n = 3. CDDP, cisplatin; CLSM, confocal laser scanning microscopy; EV, extracellular vesicle; Treg cells, regulatory T cells.

Given that TEVs can promote Treg cells’ expansion to establish an immunosuppressive tumour microenvironment, we further analysed the frequency of Treg cells among CD4+ T cells and CD8+ T cells after incubation with EVs. Based on the expression levels of CD25 and FOXP3 (markers of Treg cells) (Alonso et al. 2005; Fontenot et al. 2003), we discovered no difference in the frequency of CD8+CD25+FOXP3+ T cells in all the groups of CD8+ T cells after incubation with EVs or CDDP‐EVs, indicating few effects of these CDDP‐EVs on the differentiation of CD8+ T cells (Figure S7). In contrast to CD4+ T cells cultured alone (6.3% ± 0.1%), incubation with EVs did not significantly alter Treg levels (6.1% ± 0.3%), whereas CDDP‐EVs notably increased the frequency of CD25+FOXP3+ Treg cells to 9.6% ± 0.6%—representing a relative increase of 55.9% compared to the EV‐treated group (Figure 2f, g). These results revealed that CDDP‐EVs promote Treg cell differentiation from CD4+ T cells, rather than causing cell cytotoxicity, confirming that CDDP affected immune response through inducing TEVs to promote the amount of intratumoural Treg cells in ovarian cancers.

3.3. miR‐181a‐5p‐Enriched TEVs Promote Treg Cell Differentiation

We proceeded to investigate how CDDP‐EVs promote the differentiation of Treg cells. Previous studies have demonstrated that EVs carry abundant miRNAs that play a vital role in modulating recipient cells (Au Yeung et al. 2016; Alexander et al. 2015). To identify the specific miRNAs involved, small RNA sequencing was performed on SKOV3‐EVs and SKOV3‐CDDP‐EVs. Bioinformatic analysis of the sequencing data, including the generation of heatmaps and MA plots (Figure 3a, b), revealed a total of 68 differentially expressed miRNAs. Among these, 56 were upregulated, and 12 were downregulated in the CDDP‐EVs compared to controls. Notably, miR‐181a‐5p and miR‐92a‐3p exhibited both high expression levels and significant differential expression, prompting their selection for further investigation. Analysis of the Gene Expression Omnibus (GEO) database indicated that miR‐181a‐5p and miR‐182a‐5p are consistently highly expressed across eight ovarian cancer cell lines (Figure S8), suggesting their potential role as key regulators in ovarian cancer drug resistance. Based on these findings, miR‐181a‐5p, miR‐182a‐5p and miR‐92a‐3p were of high clinical value and thus selected for further investigation.

FIGURE 3.

FIGURE 3

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CDDP augments miR‐181a‐5p‐enriched TEVs to promote Treg cell differentiation. (a) Heatmap of differentially expressed miRNAs between SKOV3‐EVs and SKOV3‐CDDP‐EVs. Expression values are presented as log2CPM (red, high; blue, low). (b) MA plot visualising miRNA expression differences between SKOV3‐EVs and SKOV3‐CDDP‐EVs. Blue dots indicate significantly down‐regulated miRNAs, while red dots represent significantly upregulated miRNAs. (c, d) Fold change of the expressions of miR‐181a‐5p, miR‐182a‐5p and miR‐92a‐3p in SKOV3‐EVs (c) and ID8‐EVs (d) after CDDP treatment by RT‐qPCR, n = 3. (e, f) Representative flow cytometry plots (e) and quantification (f) of CD25+FOXP3+ Treg cells among CD4+ T cells. Cells were treated with NC mimic or miR‐181a‐5p, miR‐182a‐5p and miR‐92a‐3p mimics, n = 3. All statistical data were collected from three independent experiments and presented as mean values ± SD; ***p ≤ 0.001, **p ≤ 0.01, *p ≤ 0.05, ns p > 0.05 by Student's t test. CDDP, cisplatin; EV, extracellular vesicle; RT‐qPCR, reverse transcription quantitative polymerase chain reaction; TEV, tumor‐derived EV; Treg cells, regulatory T cells.

To further investigate this, the expression levels of miR‐92a‐3p, miR‐182a‐5p and miR‐181a‐5p in EVs derived from SKOV3 and ID8 cells following CDDP treatment were examined. RT‑qPCR analysis revealed that CDDP treatment significantly upregulated the expression of all three miRNAs in SKOV3‑EVs (Figure 3c). In ID8‐CDDP‐EVs, a significant increase was observed in miR‐182a‐5p and miR‐181a‐5p, while miR‐92a‐3p showed a non‐significant trend towards upregulation (Figure 3d). These results suggest that CDDP modify the molecular cargo of TEVs potentially through enriching specific miRNAs within tumour cells, which may contribute to alterations in TEV quantity and composition.

Then, to explore whether tumour‐derived miRNAs in EVs could promote Treg cell differentiation, these three upregulated and ovarian cancer‐relevant miRNA mimics (miR‐92a‐3p, miR‐182a‐5p and miR‐181a‐5p, see sequences listed in Table S1), as well as a negative control mimic (NC mimic, see sequence listed in Table S1) were then separately transfected into CD4+ T cells (Figure S9). After incubation, the frequency of Treg cells differentiated from CD4+ T cells in different groups was analysed by flow cytometry. Compared with the control group (Treg cells: 4.13%), all miRNA mimic treatments increased the proportion of Treg cells (Figure 3e). In particular, CD4+ T cells treated with the miR‐181a‐5p mimic generated the highest differentiation frequency (Treg cells: 15.8%), which was approximately 2.4‐, 1.7‐ and 1.8‐fold higher than that observed in the NC mimic (Treg cells: 6.72%), miR‐182a‐5p mimic (Treg cells: 9.49%) and miR‐92a‐3p mimic (Treg cells: 8.69%), respectively (Figure 3f). These results suggest a primary role for miR‐181a‐5p in mediating the enhancement of Treg cell differentiation.

To determine whether miR‐181a‐5p is delivered into T cells via EV internalisation, we first assessed its localisation within EVs using an RNase/protease and detergent protection assay. As shown in Figure S10, the EV membrane provided substantial protection for miR‐181a‐5p, with recovery rates of 42.5% in the EV + RNase A group and 38.7% in the EV+RNase A + Proteinase K group. The partial degradation observed in these groups may be attributed to a minor fraction of ruptured EVs and associated free miRNA during isolation. In contrast, prior disruption of EV membranes with detergent (EV+0.1% Triton X‐100 + RNase A group) led to a marked reduction in recovery to 15.8%. These results confirm that the majority of miR‐181a‐5p is encapsulated within EVs, supporting its potential for vesicle‐mediated delivery into recipient cells. Having established that miR‐181a‐5p is encapsulated in EVs, we next asked whether Treg differentiation requires EV internalisation. We employed Pitstop 2, a specific inhibitor of clathrin‐mediated endocytosis. First, we confirmed that Pitstop 2 effectively blocked EV uptake by CD4+ T cells: pre‑treatment with 15 µM Pitstop 2 significantly reduced the internalisation of PKH26‑labelled EVs (Figure S11). To assess the functional consequence, CD4+ T cells were pre‑treated with Pitstop 2, followed by co‑culture with CDDP‑EVs. Treatment with CDDP‑EVs alone increased the proportion of CD25+FOXP3+ Treg cells, promoting Treg differentiation (by 2.37 % compared with the T‑cell‑only control). However, this induction was substantially inhibited when EV internalisation was blocked by Pitstop 2 (Figure S12). These results demonstrate that EV uptake is a critical step for CDDP‑EV‐mediated Treg differentiation.

3.4. SIRT1 Is a Direct Downstream Target of miR‐181a‐5p in CD4+ T Cells

To elucidate the mechanism underlying the impact of miR‐181a‐5p on CD4+ T cells, we investigated its targets in CD4+ T cells. FOXP3 is a master regulator of transcription in Treg cells for their development and differentiation (Lu et al. 2017). Based on the Search Tool for the Retrieval of Interacting Genes (STRING) database (version 11.5), protein interactors of FOXP3 were identified under the following parameter settings: a maximum of 30 interactors to display and a minimum interaction score threshold of 0.90. A total of 30 interacting proteins were retrieved under these high‐stringency conditions. Putative target genes of miR‐181a‐5p, miR‐182a‐5p and miR‐92a‐3p were predicted using the Encyclopedia of RNA Interactomes (ENCORI) database. The intersection between the gene set of FOXP3‐interacting proteins and the predicted targets of each miRNA revealed potential candidate targets, as shown in the Venn diagrams (Figure 4a). To elucidate the regulatory network, a protein–protein interaction (PPI) analysis was performed using the STRING database between FOXP3 and the candidate target genes identified from the intersection of miRNA predictions and FOXP3‐associated genes. As shown in Figure S13, this PPI network revealed that among the potential targets of the miRNAs, the proteins SIRT1, NFATC2, JUN and FOS—predicted targets of miR‐181a‐5p—exhibited strong interactions with FOXP3. This bioinformatic analysis indicates that miR‐181a‐5p likely targets genes like SIRT1, NFATC2, JUN and FOS, which interact closely with FOXP3. This mechanism explains its strong impact on Treg cell differentiation, as seen in our cellular experiments. To validate a direct target, we first assessed the protein levels of all predicted candidates using WB assay. Notably, incubation of CD4+ T cells with the miR‐181a‐5p mimic, in contrast to the miR‐92a‐3p and miR‐182a‐5p mimic‐treated groups, led to an approximately 29% reduction in SIRT1 protein expression (Figure 4b, c). Given that previous studies have demonstrated that inhibition of SIRT1 increases the acetylation level of FOXP3, thereby promoting the induction and activation of Treg cells, these findings suggest a mechanistic link between miR‐181a‐5p and Treg differentiation via SIRT1‐FOXP3 signalling (Yang et al. 2018; Van Loosdregt et al. 2010). Furthermore, the expression level of JUN, FOS and NFATC2 protein remained largely unaltered following miR‐181a‐5p mimic treatment (Figure S14). Full, uncropped immunoblots are provided as Figure S15. Collectively, these findings provide additional evidence that miR‐181a‐5p‐enriched EVs promote CD4+ T cell differentiation primarily through suppression of SIRT1.

FIGURE 4.

FIGURE 4

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SIRT1 is a functional target of miR‐181a‐5p‐enriched EVs in CD4+ T cells. (a) Venn diagrams of the overlap between the predicted target genes of miR‐181a‐5p, miR‐92a‐3p and miR‐182a‐5p (red) and the genes associated with FOXP3 regulation (blue). (b) WB assays of SIRT1 expression in CD4+ T cells transfected with NC mimic, miR‐181a‐5p mimic, miR‐182a‐5p mimic or miR‐92a‐3p mimic. (c) Quantification of SIRT1 protein levels from (b), normalised to the NC mimic group, n = 3. (d) WB assays of SIRT1 expression in CD4+ T cells treated with CDDP‐EVs, CDDP‐EVs/mismatch or CDDP‐EVs/anti‐miR181a‐5p. (e) Quantification of SIRT1 protein levels from (d), normalised to the untreated group, n = 4. (f, g) Representative flow cytometry plots (f) and quantification (g) of CD25+FOXP3+ Treg cells among CD4+ T cells. Cells were treated with CDDP‐EVs, CDDP‐EVs/mismatch or CDDP‐EVs/anti‐miR181a‐5p, n = 3. All statistical data were collected from three independent experiments and presented as mean values ± SD; ***p ≤ 0.001, **p ≤ 0.01, *p ≤ 0.05, ns p > 0.05 by Student's t test. CDDP, cisplatin; EV, extracellular vesicle; NC, negative control; SIRT1, sirtuin 1; Treg cells, regulatory T cells; WB, western blotting.

To further investigate the on‐target effect of miR‐181a‐5p‐enriched EVs in promoting Treg cells, we designed a complementary sequence of miR‐181a‐5p and transfected it into CD4+ T cells along with CDDP‐EVs to assess the frequency of differentiating into Treg cells. Compared with the group treated with CDDP‐EVs and mismatching RNA (termed CDDP‐EVs/mismatch), the group treated with CDDP‐EVs and anti‐miR‐181a‐5p showed a considerable recovery in the expression level of SIRT1 in CD4+ T cells, verifying the potential of miR‐181a‐5p as a target for regulating SIRT1 (Figure 4d, e). Furthermore, we assessed the differentiation frequency of CD4+ T cells into Treg cells via flow cytometry. The results revealed that CDDP‐EVs significantly enhanced Treg differentiation compared to the control group (12.3% ± 1.1% vs. 4.4% ± 0.1%). However, this promotional effect was effectively abolished by anti‐miR‑181a‑5p treatment, which reduced the Treg frequency to a level comparable to that of the control (5.6% ± 0.6%) (Figure 4f, g). Thus, our findings suggest that miR‐181a‐5p is a potent candidate for enhancing the immune response in immunotherapy‐based combination therapy, which may contribute to improved clinical outcomes in ovarian cancer.

3.5. Inhibition of miR‐181a‐5p Attenuates Long‐Term CDDP‐Induced T Cell Senescence

To validate the role of miR‐181a‐5p as a detrimental factor during long‐term CDDP treatment, we established an intraperitoneal dissemination mouse model of ovarian cancer using ID8‐LUC cells, which recapitulates the high propensity for peritoneal metastasis characteristic of this disease (Figure 5a). Mice were treated every 3 days with NS, CDDP alone or CDDP combined with anti‐miR‐181a‐5p‐loaded EVs. All mice were euthanised on Day 24 post‐treatment initiation for analysis to evaluate the therapeutic effect of miR‐181a‐5p inhibition on tumour growth and metastasis. Combined treatment with CDDP and anti‐miR‐181a‐5p‐loaded EVs effectively suppressed increases in body weight and abdominal perimeter of mice. Inhibition rates reached 13.6% and 14.2% compared to the CDDP‐alone group, and 19.0% and 21.9% compared to the NS group, respectively (Figure 5b, c). The reductions in body weight and abdominal perimeter indicate that CDDP treatment effectively attenuated tumour progression, and the addition of anti‐miR‐181a‐5p‐loaded EVs enhanced this therapeutic efficacy. Bioluminescence imaging via IVIS on Day 24 revealed a significant reduction in abdominal fluorescence across all treatment groups compared to the NS group. Notably, the combined treatment of CDDP and anti‐miR‐181a‐5p‐loaded EVs exhibited the lowest average fluorescence intensity among all groups (Figure 5d, e), indicating that the incorporation of anti‐miR‐181a‐5p‐loaded EVs potently enhanced the suppression of both tumour growth and metastasis. Macroscopic examination revealed varying degrees of tumour dissemination, visible as white metastatic foci, on the abdominal wall and mesentery in all groups (indicated by blue coils; Figure 5f). In addition, both the NS and CDDP groups exhibited hemorrhagic ascites, whereas the combined treatment of CDDP and anti‐miR‐181a‐5p‐loaded EVs showed ascitic fluid with markedly lighter colouration. All macroscopic tumour nodules larger than 1 mm in diameter were excised, photographed, and quantified. A significant reduction in tumour number was observed in the combined treatment group compared to the other two groups (Figure 5g, h). Collectively, targeted inhibition of miR‐181a‐5p effectively suppresses CDDP resistance and attenuates ovarian cancer progression in vivo. The level of miR‐181a‐5p in EVs isolated from ascites collected on Day 0 and Day 24 was measured. On Day 0, the abundance of miR‐181a‐5p was comparable between the CDDP group and the combined treatment group. However, on Day 24, the latter showed a significantly lower level than the former, indicating effective silencing of miR‐181a‐5p by the anti‐miR‐181a‐5p construct (Figure 5i). Immunofluorescence staining of tumour nodule sections for Foxp3 revealed the strongest and most extensive fluorescence signal in the CDDP group. The signal in the CDDP in combination with anti‐miR‐181a‐5p‐loaded EVs was slightly higher than that in the NS group, though the difference was not remarkable (Figure 5j, k). CDDP treatment led to a marked increase in Foxp3+ cells, most likely Treg cells, in the tumour microenvironment—an effect that was partially mitigated by EV‐mediated inhibition of miR‐181a‐5p. This increase suggests a potential association with T cell senescence or enhanced Treg induction, a possibility that warrants further investigation. To confirm whether the mechanism by which miR‐181a‐5p influences T cells differentiation via SIRT1 suppression operates in vivo, we performed multiplex immunofluorescence staining on tumour tissue (CD4: FITC, green; SIRT1: Cy5, red; nuclei: DAPI). As shown in Figure S16, CDDP monotherapy significantly reduced SIRT1 expression in CD4+ T cells compared with the NS group, whereas this reduction was markedly reversed in the group receiving CDDP combined with anti‐miR‐181a‐5p‐loaded EVs. These findings confirm that miR‐181a‐5p can modulate T cells differentiation by downregulating the SIRT1 pathway, consistent with the conclusions drawn from cellular experiments.

FIGURE 5.

FIGURE 5

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Inhibition of miR‐181a‐5p enhances the efficacy of CDDP in vivo. (a) Schematic diagram of therapeutic investigation in the intraperitoneal tumour‐bearing mice ovarian cancer model in vivo. Mice were randomised into three groups: (i) NS, (ii) CDDP alone and (iii) CDDP + EVs/anti‐miR‐181a‐5p. (b) Body weight of mice, n = 3. (c) Abdominal perimeter of mice, n = 3. (d, e) Representative bioluminescence imagings (d) and quantitative analysis (e) of mice on Day 24, n = 3. (f) Representative gross images of abdominal metastases observed during terminal dissection. (g, h) Images (g) and total numbers (h) of excised tumours (diameter > 1 mm) on Day 24, n = 3. (i) Relative expression levels of miR‐181a‐5p in ascites samples collected on Day 0 and Day 24, as determined by RT‐qPCR, n = 3. (j) Immunofluorescence imaging of Foxp3 expression (PE) in tumour tissues. Nuclei are counterstained with DAPI. Scale bar, 50 µm. (k) Quantification of the mean fluorescence intensity of Foxp3 from each treatment group, n = 3. All statistical data were collected from three independent experiments and presented as mean values ± SD; ***p ≤ 0.001, **p ≤ 0.01, *p ≤ 0.05, ns p > 0.05 by Student's t test. CDDP, cisplatin; EV, extracellular vesicle; NS, normal saline; RT‐qPCR, reverse transcription quantitative polymerase chain reaction.

To validate that miR‐181a‐5p inhibition potentiates CD4+/CD8+ T cell‐mediated immunity and attenuates Treg suppression, we performed flow cytometric analysis of T cell infiltration in tumours and spleens. As shown in Figure 6a–c, analysis of splenocytes revealed that the proportion of CD3+CD4+ T cells in the CDDP in combination with anti‐miR‐181a‐5p‐loaded EVs group (1.31% ± 0.50%) was 4.23‐ and 2.52‐fold higher than that in the NS and CDDP groups, respectively. Similarly, the proportion of CD3+CD8+ T cells (6.69% ± 1.58%) was 2.3‐ and 1.26‐fold greater than that in the NS and CDDP groups, respectively. A similar trend was observed in tumour‐infiltrating lymphocytes. The combination therapy group exhibited a higher proportion of T cells than the NS and CDDP groups: CD3+CD4+ T cells (25.87% ± 3.02%) increased by 2.4‐ and 1.55‐fold, and CD3+CD8+ T cells (25.13% ± 2.94%) increased by 3.7‐ and 1.59‐fold, respectively (Figure 6d–f). The frequency of CD25+Foxp3+ Treg cells in ascitic fluid was evaluated on Days 0, 9 and 24. No statistically significant differences in Treg frequency were observed among the groups on either Day 0 or Day 9 (Figure 6g¸h and j¸k). However, a non‐significant decrease in Treg frequency was noted in the combination therapy group relative to both the NS and CDDP groups on Day 9. The frequency of immunosuppressive Treg cells measured on Day 24 was significantly lower in the combination therapy group (6.83% ± 1.20%) than in both the CDDP group (21.07% ± 6.37%) and the NS group (69.47% ± 5.61%), indicating the minimal Treg infiltration achieved by the combination therapy (Figure 6i, l). In summary, the delivery of anti‐miR‐181a‐5p via EVs significantly enhanced the response to CDDP treatment. This effect was mediated through the silencing of miR‐181a‐5p, which alleviated immunosuppression by reducing the infiltration of Treg cells into ascites and promoting the activation and proliferation of CD4+ and CD8+ T cells, thereby potentiating antitumour immunity and ultimately attenuating CDDP resistance.

FIGURE 6.

FIGURE 6

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Inhibition of miR‐181a‐5p reprograms antitumour immunity by promoting T cell infiltration and suppressing immunosuppression. (a–c) Representative flow cytometry plots (a) and quantification (b, c) of the percentage of CD3+CD4+ T cells and CD3+CD8a+ T cells in spleen by flow cytometry, n = 3. (d–f) Representative flow cytometry plots (d) and quantification (e, f) of the percentage of CD3+CD4+ T cells and CD3+CD8a+ T cells in tumour by flow cytometry, n = 3. (g–l) Representative flow cytometry plots (g–i) and quantification (j–l) of CD25+Foxp3+ Treg cells in ascites samples collected on Days 0, 9 and 24 by flow cytometry, n = 3. All statistical data were collected from three independent experiments and presented as mean values ± SD; ***p ≤ 0.001, **p ≤ 0.01, *p ≤ 0.05, ns p > 0.05 by Student's t test.

4. Disscussion

Characterized by high rates of relapse and metastasis, ovarian cancer accounts for the highest mortality among gynaecological malignancies, calling for the study of innovative treatment strategies for prolonged overall survival (Yap et al. 2009; Ahmed and Stenvers 2013). Considering the immunogenic microenvironment of ovarian cancer, which is often characterised by lymphocyte infiltration, immunotherapy by eliciting or enhancing a synergistic immune response is considered an effective treatment strategy. CDDP, the most widely used clinical treatment for ovarian cancer, has recently been evaluated in combination with immunotherapy (Konstantinopoulos and Cannistra 2021). However, first‐line clinical trials of such combinations have not yielded significant survival benefits. This limited efficacy suggests that CDDP may exert complex immunomodulatory effects within the tumour microenvironment, which could either support its antitumour activity or paradoxically contribute to treatment resistance (Meng et al. 2024). Notably, although CDDP is known to alter immune cell function, the precise mechanisms underlying its systemic immunomodulatory effects remain unclear.

Carrying an assortment of proteins, lipids and miRNAs, EVs serve as mediators in intercellular communication (Maia et al. 2018). Previous studies have demonstrated that TEVs‐mediated communication between cancer cells and immune cells plays a key role in regulating immune response (Whiteside 2016). In particular, many miRNAs carried by TEVs were found to disable the efficacy of immune cells (Au Yeung et al. 2016; Alexander et al. 2015). Recent studies have increasingly highlighted the role of EVs as critical mediators of chemoresistance and tumour microenvironment remodelling. In high‑grade serous ovarian cancer, EVs have been shown to carry the lncRNA CATED, which activates the MAPK pathway and contributes to platinum resistance (Liu et al. 2025). Additionally, CDDP‑stimulated ovarian cancer stem cells release EVs that can reprogram mesenchymal stem cells towards a pro‑tumourigenic phenotype (Vera et al. 2019). Building on this evidence, we hypothesised that CDDP might similarly reprogram the cargo of tumour‑derived EVs, and that these modified EVs could function as a key intermediary in transmitting immunomodulatory signals from cancer cells to T cells within the tumour microenvironment.

Using the widely used platinum agent CDDP as a model, we investigate d CDDP‐induced EVs from ovarian cancer cells and found that CDDP significantly promote d the enrichment of miR‐181a‐5p within these EVs, which subsequently enhance d the differentiation of CD4+ T cells towards Treg cells through inhibiting SIRT1 expression. Specifically, our results first show ed that CDDP mediate d a dramatic increase in the secretion of EVs from ovarian cancer cells. In contrast, CDDP barely affect ed the secretion of EVs from non‐cancerous cells. Furthermore, CDDP‐induced TEVs were highly enriched with miR‐181a‐5p, which promote d Treg cell differentiation by targeting SIRT1 to stimulate FOXP3 expression. In tumour‐bearing mice, treatment with anti‐miR‐181a‐5p‐loaded EVs reduce d Treg infiltration, enhance d CD4+ and CD8+ T cell‐mediated antitumour immunity, and significantly restrain ed tumour growth. Collectively, these findings uncover a key mechanism through which CDDP exacerbates Treg‐mediated immunosuppression. Therapeutic targeting of this pathway offers a promising approach to augment combination immunotherapy in ovarian cancer and potentially improve patient survival.

The functional significance of miR‑181a‑5p extends beyond immune regulation, as it broadly influences tumour progression across multiple cancer types. For instance, miR‑181a‑5p promotes resistance to radioactive iodine therapy in thyroid cancer by downregulating the iodide transporter NIS (Gierlikowski et al. 2021), and drives gastric cancer progression through repression of tumour suppressors MEG2 (Liu et al. 2017). Moreover, EV‑derived miR‑181a‑5p in serum serves as a diagnostic biomarker for bone‑metastatic prostate cancer, highlighting its role in metastatic dissemination (Wang et al. 2021). These findings, together with our current study revealing its function in promoting Treg‑mediated immunosuppression via the SIRT1/FOXP3 pathway, establish miR‑181a‑5p as a central regulator that concurrently coordinates tumour‑intrinsic progression and immune evasion, positioning it as a promising therapeutic target across multiple cancers.

However, this study has several limitations. First, the findings are primarily derived from cell lines and murine models. Validating the enrichment of miR‐181a‐5p in tumour‐derived EVs from ovarian cancer patient samples and correlating these levels with Treg infiltration and clinical outcomes represent critical steps towards clinical translation. Second, while this study focused on miR‐181a‐5p, the immunomodulatory effects observed may also be influenced by additional bioactive cargo within CDDP‐EVs, whose contributions warrant systematic investigation in future studies. Finally, although our results demonstrate therapeutic promise in murine models, the safety profile, optimal dosing and long‐term effects of EV‐based delivery systems require comprehensive preclinical evaluation before clinical application.

Author Contributions

Liyan Zheng: formal analysis (equal), methodology (equal), supervision (equal), visualisation (equal), writing – original draft (equal), writing – review and editing (equal). Sijia Mu: data curation (lead), investigation (equal), validation (equal), visualisation (lead), software (equal), writing – original draft (equal). Mingxue Liu: data curation (lead), investigation (equal), validation (equal), software (equal). Panmin Lei: data curation (supporting), investigation (equal), validation (equal). Jingchen Zhang: data curation (supporting), investigation (equal), validation (equal). Yuting Dai: data curation (supporting), software (equal), visualisation (supporting). Chenxi He: investigation (equal), validation (equal). Fengli Qu: conceptualisation (equal), funding acquisition (lead), resources (supporting), project administration (equal), supervision (equal), writing – review and editing (equal). Weihong Tan: conceptualisation (equal), resources (supporting), writing – review and editing (equal).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting Information: jev270261‐sup‐0001‐SuppMat.docx

JEV2-15-e70261-s001.docx (16.1MB, docx)

Acknowledgements

This work was funded by the National Natural Science Foundation of China (Nos. U25A20561, 22504154, 22274141), Zhejiang Provincial Health Commission Research Foundation (WKJ‐ZJ‐242), the foundation of Hangzhou Institute of Medicine, CAS (2024ZZBS04, 2024ZZBS08), the Leading Innovative and Entrepreneur Team Introduction Program of Zhejiang Province (2024R01005), the Pioneer Research and Development Program of Zhejiang (2023SDYXS0002, 2025SDYXS0001) and Research Funds of Hangzhou Institute for Advanced Study, UCAS (2023HIAS‐Y029).

Data Availability Statement

The data that supports the findings of this study are available in the supplementary material of this article.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Information: jev270261‐sup‐0001‐SuppMat.docx

JEV2-15-e70261-s001.docx (16.1MB, docx)

Data Availability Statement

The data that supports the findings of this study are available in the supplementary material of this article.

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