An Engineered Extracellular Vesicle With Enhanced Tumor and Lymph Nodes Targeting as siRNA Delivery System for Robust Tumor Immunotherapy
Yusi Wang, Rui Zhang, Xuejing Zhou, Lin Tang, Die Hu, Yibing Zhang, Yuling Yang, Bailing Zhou, Li Yang
TL;DR
Engineered extracellular vesicles with enhanced targeting abilities deliver siRNA to boost tumor immunotherapy and prolong survival in mice.
Contribution
A novel EV platform with DP7-C peptide engineering enables dual targeting to tumors and lymph nodes for effective immunotherapy.
Findings
DP-EVs show enhanced accumulation in lymph nodes and tumor tissues.
DP-EVs/siPD-L1 inhibits tumor growth and prolongs survival in mice.
The therapy activates immune responses and remodels the tumor microenvironment.
Abstract
Tumor‐derived extracellular vesicles (EVs) are a class of natural nanocarriers with phospholipid bilayers that show great promise as personalized cancer vaccine platforms due to their ability to carry tumor‐specific antigens. However, their immunotherapeutic potential is hindered by limited tissue‐specific targeting. In this study, we engineered tumor cell‐derived EVs using an immunomodulatory peptide, DP7‐C, to generate DP7‐C engineered EVs (DP‐EVs). These DP‐EVs exhibited significantly enhanced accumulation in both lymph nodes and tumor tissues. Additionally, they demonstrated improved cellular uptake and facilitated more efficient endosomal escape. To further enhance the therapeutic efficacy, programmed cell death 1 ligand 1 targeting small interfering RNA (siPD‐L1) was loaded into the DP‐EVs, resulting in DP‐EVs/siPD‐L1. This formulation enabled concurrent suppression of PD‐L1…
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FIGURE 7- —National Key Research and Development Program of China10.13039/501100012166
- —National Natural Science Foundation of China10.13039/501100001809
- —1.3.5 project for disciplines of excellence, West China Hospital, Sichuan University
- —Sichuan Science and Technology Program
- —National Clinical Research Center for Geriatrics, West China Hospital, Sichuan University
- —The Frontiers Medical Center, Tianfu Jincheng Laboratory Foundation
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Taxonomy
TopicsExtracellular vesicles in disease · Nanoplatforms for cancer theranostics · interferon and immune responses
Introduction
1
Cancer immunotherapy, which harnesses the body's immune system to combat malignancies, has revolutionized oncology by eliciting durable clinical responses in various cancers [1, 2]. Among these strategies, cancer vaccines hold considerable promise for their potential to induce tumor‐specific immune memory [3]. However, the clinical efficacy of traditional vaccine platforms, including those based on whole cells, tumor‐associated antigens (TAAs), or peptides, is often limited by insufficient immunogenicity and inadequate delivery to immune cells, failing to achieve robust antitumor immunity [4, 5].
EVs, particularly those derived from tumor cells, have emerged as a promising alternative platform for next‐generation cancer vaccines. As natural lipid bilayer nanoparticles with a diameter ranging from 30 to 200 nm, EVs are enriched with source cell‐specific components, serving as messengers for intercellular communication [6, 7, 8, 9, 10]. Notably, EVs secreted by tumor cells are enriched with unique tumor antigens, positioning them as ideal candidates for personalized vaccines [11, 12, 13]. However, the intrinsic complexity of cancer biology, coupled with the immunosuppressive TME poses significant challenges to the effectiveness of EV‐based therapies [14]. Combining vaccines with immune checkpoint blockade (ICB) strategies, such as targeting the cytotoxic T‐lymphocyte‐associated protein 4 (CTLA‐4) and programmed cell death 1/programmed cell death 1 ligand 1 (PD‐1/PD‐L1) pathway, has emerged as a promising approach to overcoming immunosuppression in the TME [15, 16]. However, the systemic delivery of ICB agents often results in off‐target effects and dose‐limiting toxicities, highlighting an urgent need for strategies that enable precise delivery of immunomodulatory agents to key immune organs [17, 18].
siRNA offers a potential solution by enabling precise silencing of immunosuppressive molecules like PD‐L1 within target cells [19, 20]. However, the clinical translation of siRNA is hindered by its instability and poor cellular uptake, necessitating the development of effective delivery vehicles [21]. As naturally derived nanoparticles, EVs possess several inherent advantages for siRNA delivery, including superior biocompatibility, low immunogenicity, and innate targeting capabilities [22, 23, 24]. Despite these advantages, the native targeting efficiency of EVs is often insufficient for specific therapeutic applications, emphasizing the need to engineer EVs with enhanced targeting capabilities [25].
In this study, we developed an engineered EV‐based vaccine to address these challenges in a synergistic manner. We functionalized tumor cell‐derived EVs with cholesterol‐modified DP7 (DP7‐C), a peptide with dual functionality. DP7 (VQWRIRVAVIRK) is an antimicrobial peptide, and after cholesterol modification, DP7‐C functions both as a delivery carrier and an immune adjuvant, as demonstrated in our previous research [26, 27, 28, 29]. Briefly, as a carrier, DP7‐C can efficiently deliver siRNA and peptides to target cells and tissues without causing significant toxicity or side effects [28, 30]. As an immune adjuvant, DP7‐C activates the MyD88–IKK–IκB–NF‐κB signaling pathway in BMDCs, significantly upregulating MyD88 and stimulating DC maturation and the release of proinflammatory cytokines [28]. In comparison with EVs derived from untreated tumor cells, DP‐EVs exhibited enhanced accumulation in lymph nodes and tumor tissues, improved cellular uptake, and superior endosomal escape ability. Furthermore, we loaded siPD‐L1 into DP‐EVs, creating a dual‐function complex: DP‐EVs/siPD‐L1. This complex is designed to simultaneously deliver tumor antigens and silence PD‐L1 expression in both DCs within lymph nodes and tumor cells, thereby reversing local immunosuppression and amplifying vaccine‐elicited immune responses [31, 32, 33, 34].
Our results demonstrate that DP‐EVs/siPD‐L1 exhibit potent efficiency in the knockdown of target genes in both in vitro and in vivo settings. Moreover, this complex significantly inhibited tumor growth and prolonged survival in tumor‐bearing mice by potently activating immune cells in the lymph nodes and remodeling the immunosuppressive TME. This work presents a novel EV‐based vaccine strategy that integrates targeted delivery with siRNA‐mediated ICB, offering a promising and translatable approach for cancer immunotherapy.
Results
2
Preparation and Characterization of EVs and DP‐EVs
2.1
Prior to constructing DP‐EVs, we first evaluated the potential cytotoxicity of DP7‐C toward E.G7‐OVA cells. Results showed that treatment with 50 µg/mL DP7‐C did not affect cell viability compared with untreated controls (Figure S1A), indicating that this concentration is biocompatible and suitable for subsequent stimulation experiments. EVs and DP‐EVs were then isolated from cell culture supernatants via ultracentrifugation (Figure 1A) and characterized using a panel of standard techniques. Western blot analysis confirmed the presence of canonical EV markers (CD9, CD63, TSG‐101) in both EVs and DP‐EVs (Figure 1B), verifying the successful isolation of vesicular structures. Transmission electron microscopy (TEM) imaging revealed their typical cup‐shaped morphology (Figure 1C), while nanoparticle tracking analysis (NTA) showed that DP‐EVs exhibited a slightly larger average particle size (164.0 nm) than EVs (153.9 nm) (Figure 1D). In contrast, the zeta potentials of DP‐EVs (−40.77 ± 0.44 mV) and EVs (−40.99 ± 0.80 mV) were almost identical (Figure 1E). Together, these data confirm that the fundamental morphological and structural integrity of EVs is preserved following DP7‐C engineering.
Generation and characterization of EVs and DP‐EVs. (A) Schematic illustration of the preparation of EVs and DP‐EVs. (B) Western blot analysis of EV marker proteins (CD9, CD63, and TSG‐101). (C) TEM images showing the morphology of EVs and DP‐EVs. Scale bars, 100 nm. (D) Size distribution of EVs and DP‐EVs measured by NTA. (E) Zeta potential of EVs and DP‐EVs determined by NTA. (F–H) Volcano plot, KEGG, and GO analyses of proteomic sequencing in EVs and DP‐EVs. (I–K) Volcano plot, KEGG, and GO analyses of miRNA sequencing in EVs and DP‐EVs. N = 3.
To further investigate the effect of DP7‐C incorporation during EV biogenesis, we performed proteomic and microRNA (miRNA) profiling of EVs and DP‐EVs. Volcano plots revealed pronounced differences in molecular composition, with 124 proteins upregulated and 86 downregulated in DP‐EVs compared with native EVs (Figure 1F). Likewise, miRNA array analysis identified four upregulated and 10 downregulated miRNAs (Figure 1G), indicating that DP7‐C stimulation not only influences vesicle protein content but also modulates their RNA cargo. Functional enrichment analyses provided mechanistic insights into these alterations. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis of the differentially expressed proteins (Figure 1H) and miRNAs (Figure 1I) revealed significant enrichment in pathways associated with Fc gamma R‐mediated phagocytosis, regulation of the actin cytoskeleton, and endocytosis. These pathways are closely related to vesicle–cell interactions and uptake processes, suggesting that DP7‐C modification may facilitate enhanced EV internalization by recipient cells. This observation is consistent with a previous finding, which reported that the elevated endocytosis and actin cytoskeleton regulatory pathways contributed to the increased phagocytic efficiency of 3D‐Exo by osteoblasts [35]. A Gene Ontology (GO) analysis also suggested alterations in biological processes including vesicle‐mediated transport and the regulation of the MAPK and JNK cascades (Figure 1J,K), which are known to participate in immune activation and stress signaling. Collectively, these omics findings strongly suggest that DP7‐C engineering actively reprograms the molecular cargo of EVs, potentially endowing them with enhanced functional capabilities relevant to immune cell interactions and intracellular trafficking.
DP‐EVs Exhibit Enhanced Lymph Node and Tumor‐Targeting Capacity
2.2
To systematically evaluate the targeting specificity of engineered EVs, DIR‐labeled naive EVs and DP‐EVs were subcutaneously administered to mice bearing E.G7‐OVA tumors (Figure 2A). In vivo imaging and quantitative analysis confirmed the accumulation of both vesicle types in tumor tissues and draining lymph nodes, as well as their presence in the liver, consistent with previous reports [22, 23, 24] (Figures 2B,C and S1B). However, DP‐EVs displayed a markedly higher fluorescence intensity in both lymph nodes and tumor tissues. This observation was further corroborated by ex vivo imaging of frozen tissue sections (Figure 2D). To determine whether this targeting enhancement was specific to the E.G7‐OVA model, we next evaluated 4T1‐derived EVs and DP‐EVs in a parallel experiment. Consistent with our previous findings, 4T1‐derived DP‐EVs also exhibited significantly enhanced accumulation in tumors and lymph nodes compared with EVs (Figure S2A,B). This reproducibility across distinct tumor models indicates that the improved targeting conferred by DP7‐C modification is a generalizable property, not limited to a particular cell lineage.
*DP‐EVs exhibit enhanced accumulation in tumors and lymph nodes. (A) Experimental timeline of in vivo biodistribution. (B and C) Fluorescence imaging of normal saline (NS group), DIR‐labeled EVs and DP‐EVs 24 h postinjection in E.G7‐OVA tumor bearing mice. (D) Representative CLSM images of frozen sections of major organs after subcutaneous injection of DIR‐EVs and DIR‐DP‐EVs. Scale bars, 100 µm. (E and F) Flow cytometry of LNs and tumor after subcutaneous injection of DIR‐EVs and DIR‐DP‐EVs in E.G7‐OVA tumor bearing mice. (G) Schematic illustration of the LNs and tumor targeting mechanism for DP‐EVs. All values presented in this figure are expressed as the mean ± s.d. Student's t‐test was utilized for two‐group comparisons. *p < 0.05, *p < 0.01. N = 3.
To elucidate the mechanisms underlying the enhanced targeting, we performed flow cytometry analysis on single‐cell suspensions from lymph nodes and tumors. In lymph nodes, a significantly higher proportion of DIR‐positive cells was detected within the CD45^+^ immune cell population in the DP‐EV group compared with the EV group, whereas no notable difference was observed among nonimmune (CD45^−^) cells. Further immunophenotyping revealed that this increase was primarily associated with a elevated frequency of migratory DCs (CD11c^+^ CD103^+^) among the DIR^+^ immune cells (Figure 2E). Conversely, within tumor tissues, the enhancement in DIR^+^ cells were predominantly observed in the nonimmune cell compartment, with no significant change detected in the immune cell fraction. Notably, an increased presence of migratory DCs was also identified within the DIR^+^ immune cell subset in tumors (Figure 2F). Taken together, these findings demonstrate that subcutaneously administered EVs and DP‐EVs can be internalized by DCs, which subsequently migrate to lymph nodes and tumor sites. DP7‐C modification not only enhances this immune‐cell‐mediated trafficking but also promotes direct tumor homing, thereby enabling DP‐EVs to achieve a dual‐targeting delivery pattern (Figure 2G). This unique biodistribution profile provides a mechanistic basis for the superior immunotherapeutic performance of DP‐EVs in subsequent experiments.
DP‐EV Possess Enhanced Cellular Uptake and Endosomal Escape of EVs Through Diversified Endocytic Pathways
2.3
To investigate the cellular uptake behavior of EVs and DP‐EVs, sterile vesicles were prepared for the following studies (Figure 3A). Flow cytometric analysis revealed that DP‐EVs exhibited significantly enhanced internalization by their parental tumor cells (E.G7‐OVA), whereas uptake by bone marrow‐derived DCs (BMDCs) and macrophages (BMDMs) remained comparable to that of unmodified EVs (Figure 3B). These findings suggest that DP7‐C modification selectively promotes homologous uptake by donor cells, consistent with its role in reinforcing tumor‐specific recognition. To further delineate the cellular uptake pathways, we employed fluorescent probe colocalization assays after 24 h of incubation. Native EVs mainly colocalized with Dextran Texas and cholera toxin subunit B (CT‐B), markers of macropinocytosis and caveolin‐mediated endocytosis, respectively (Figure 3C). Notably, DP‐EVs displayed additional colocalization with transferrin, indicative of clathrin‐mediated internalization (Figure 3C). These observations were corroborated by pharmacological inhibition studies. Treatment with amiloride (macropinocytosis inhibitor) or genistein (caveolin inhibitor) reduced EV uptake, confirming that these two routes dominate for native EVs (Figure 3D). Notably, chlorpromazine (a clathrin pathway inhibitor) selectively suppressed DP‐EV uptake without affecting EV internalization, supporting the notion that DP7‐C modification introduces an additional clathrin‐dependent endocytic route (Figure 3D). Collectively, these results demonstrate that while native EVs enter BMDCs primarily through macropinocytosis and caveolin‐mediated pathways, DP‐EVs acquire enhanced versatility in uptake mechanisms, allowing more efficient cellular entry through multiple routes.
*DP‐EVs can be effectively internalized by multiple cells and facilitated BMDCs migration and maturation. (A) Schematic illustration of DP‐EVs endocytosed by BMDCs. (B) Uptake efficiency of DIO‐EVs and DP‐EVs in E.G7‐OVA, BMDCs, and BMDMs. Scale bars, 100 µm. (C) Representative colocalization images in BMDCs of three uptake pathway markers with EVs and DP‐EVs, CT‐B, a marker of the caveolin pathway; transferrin, a marker of the clathrin pathway; and Dextran Texas, a marker of the macropinocytosis pathway. Scale bars, 20 µm. (D) Uptake efficiency of DIO‐EVs and DIO‐DP‐EVs in BMDCs pretreated with three inhibitors, amiloride (20 µM, an inhibitor of the macropinocytosis pathway); chlorpromazine (5 µM, an inhibitor of the clathrin‐mediated endocytosis pathway); and genistein (30 µM, an inhibitor of the caveolin‐mediated endocytosis pathway). (E) Representative colocalization images at 24 h after incubation in BMDCs of early endosomes, late endosomes, and lysosomes with EVs and DP‐EVs. Scale bars, 10 µm. (F) The percentages of mature DCs after different treatments. (G) The proportion of M1/M2 after different treatment. (H) The expression of CCR7 after different treatments detected by qPCR. (I) The expression of CCR7, AKT, p‐AKT, PI3K, p‐PI3K, JNK, p‐JNK, MAPK, p‐MAPK, ERK, p‐ERK, p65, and p‐p65 proteins in BMDCs after different treatment. (J) Schematic illustration of DC migration caused by DP‐EVs. (K) The percentages of migrated DCs after different treatments. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001. N = 3.
Efficient cytosolic delivery of vesicle cargo depends not only on uptake but also on successful endosomal escape, as vesicles entrapped in lysosomes are rapidly degraded. To assess this process, we examined the intracellular trafficking of EVs and DP‐EVs at different time points postinternalization. Following a 6‐h period postinternalization, both EVs and DP‐EVs were observed to colocalize with late endosomes and lysosomes (Figure S3A,B). It is noteworthy that after 24 h, the colocalization of DP‐EVs with early endosomes, late endosomes, and lysosomes was significantly reduced in comparison with that of EVs (Figures 3E and S3A,B). These findings indicate that DP‐EVs possess an enhanced endosomal escape capacity.
DP‐EVs Promote BMDC Maturation and Migration via CCR7 Upregulation
2.4
To examine how EVs and DP‐EVs influence the immunological behavior of DCs, we investigated their effects on the maturation and migration of BMDCs. Flow cytometry demonstrated that both EVs and DP‐EVs promoted BMDCs maturation after 24 h of incubation; however, no significant difference was observed between the two groups (Figure 3F). Besides, results showed that DP‐EVs promoted BMDCs migration compared with EVs (Figure 3K). To explore the molecular basis of this enhanced mobility, we assessed the expression of CCR7, a chemokine receptor essential for DC homing to lymph nodes. Quantitative PCR (qPCR) analysis (Figure 3H), flow cytometry (Figure S4A), and immunofluorescence staining (Figure S4B) revealed a pronounced upregulation of CCR7 in DP‐EV‐treated BMDCs relative to EV‐treated cells, which was further confirmed by western blotting (Figure 3I). These findings demonstrate that DP‐EVs trigger CCR7‐dependent signaling to facilitate DC migration. Mechanistically, DP‐EVs activated multiple intracellular signaling pathways associated with DC motility and chemotaxis. Western blot analysis revealed increased phosphorylation of key components in the PI3K–AKT, MAPK–JNK, MAPK–ERK, and MAPK–NF‐κB cascades following 24 h of DP‐EV exposure (Figure 3I). These pathways are well known to converge on CCR7 transcriptional regulation and cytoskeletal remodeling, both of which are crucial for the directional migration of DCs toward CCL19/CCL21 gradients (Figure 3J) [36]. Taken together, these results suggest that DP7‐C engineering reprograms EV–DC interactions at the signaling level: while maintaining comparable maturation‐inducing capability, DP‐EVs more effectively activate CCR7‐associated migration pathways through coordinated PI3K and MAPK signaling. This enhanced migratory phenotype provides a plausible explanation for the superior lymph node accumulation and immunological activation observed with DP‐EVs in vivo.
DP‐EVs Can Change the Polarization of BMDMs
2.5
To investigate the impact of EVs and DP‐EVs on BMDMs, a flow cytometry analysis was conducted in order to evaluate macrophage polarization. Compared with native EVs, DP‐EVs markedly increased the ratio of M1 (CD86^+^CD206^−^) to M2 (CD86^−^CD206^+^) macrophages, indicating a pronounced shift toward a proinflammatory M1 phenotype (Figures 3G and S5A,B). To further characterize this phenotypic transition, we examined the transcriptional profile of macrophage activation markers and cytokines using qPCR. Consistent with the flow cytometry results, DP‐EV treatment significantly upregulated CD86, MHC‐I, MHC‐II, CCR7, CXCL9, CXCL10, and IL‐23α (Figure S5C). These molecules are classical hallmarks of M1‐type macrophage activation and are closely associated with antigen presentation, Th1 cell recruitment, and IFN‐γ‐driven immune responses (Figure S5). Together, these results demonstrate that DP‐EVs effectively reprogram tumor‐associated macrophages toward an M1‐like phenotype with enhanced antitumor function.
DP‐EVs Are Capable of Delivering siPD‐L1 With High Efficiency
2.6
Given the enhanced lymph node and tumor targeting as well as superior endosomal escape capability of DP‐EVs, we next explored their potential as carriers for siRNA. siPD‐L1 was loaded into both EVs and DP‐EVs using an electroporation approach (Figure 4A). Subsequently, TEM confirmed that siRNA‐loaded vesicles preserved the characteristic cup‐shaped morphology of DP‐EVs and EVs (Figures 4B and S6A). NTA data showed a modest increase in particle diameter from 168.1 to 173.8 nm for DP‐EVs and from 155.6 to 175.4 nm for EVs after siRNA loading (Figures 4C and S6B). In parallel, the zeta potential shifted toward less negative values (from –46.75 ± 0.56 to –36.85 ± 1.07 mV for DP‐EVs, and from –58.05 ± 1.06 to –29.52 ± 1.57 mV for EVs) (Figures 4D and S6C). These subtle changes confirm the successful incorporation of siRNA without disrupting vesicle integrity.
*DP‐EVs delivered siRNA to downregulate PD‐L1 both in E.G7‐OVA and BMDCs. (A) Schematic illustration of DP‐EVs loading siRNA. Scale bars = 100 nm. (B) TEM images DP‐EVs and DP‐EVs/siRNA. (C) Size detection by NTA of DP‐EVs and DP‐EVs/siRNA. (D) Zeta potential detection through NTA of DP‐EVs and DP‐EVs/siRNA. (E) RNase A protection assay of EVs/siRNA and DP‐EVs/siRNA. (F) Uptake efficiency of DIO‐EVs/cy3‐siRNA and DIO‐DP‐EVs/cy3‐siRNA. (G) Representative colocalization images at 6 and 24 h after incubation in BMDCs of early endosomes, late endosomes, and lysosomes with DP‐EVs/cy3‐siRNA. (H) Detection of PD‐L1 expression at the mRNA level in E.G7‐OVA and BMDCs by qPCR. (I and J) Detection of PD‐L1 expression at the protein level in E.G7‐OVA and BMDCs by western blot. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. ***p < 0.001, ***p < 0.0001. N = 3.
The encapsulation efficiency (EE) of siRNA was detected to be 18.93% for EVs and 21.39% for DP‐EVs (Figure S7A). To evaluate the ability the stability of the encapsulated siRNA, formulations were exposed to RNase A digestion. While free siRNA was completely degraded within 30 min, siRNA encapsulated in both EVs and DP‐EVs remained intact for up to 4 h (Figure 4E). These results confirm that DP‐EVs efficiently encapsulate and protect siRNA molecules from enzymatic degradation, maintaining structural integrity suitable for subsequent delivery studies.
DP‐EVs/siPD‐L1 Enable Efficient Gene Silencing
2.7
To assess the siRNA delivery capability of EVs and DP‐EVs, we conducted confocal microscopy (Figure S7B) and flow cytometry analyses (Figure 4F). Whereas free siRNA exhibits minimal cellular uptake, it can be effectively internalized by BMDCs and E.G7‐OVA when delivered via EV or DP‐EV. Notably, DP‐EVs exhibited significantly higher intracellular delivery efficiency in E.G7‐OVA compared with native EVs. Consistent with a previous literature report [37], free siRNA is incapable of efficient endosomal escape within 24 h postcellular entry (Figure S8A), necessitating the use of delivery vectors for functional gene silencing. Endosomal trafficking analysis showed that siRNA delivered via DP‐EVs exhibited markedly reduced colocalization with early endosomes, late endosomes, and lysosomes after 24 h, indicating improved cytosolic release (Figures 4G and S8B).
To evaluate functional gene silencing, E.G7‐OVA and BMDCs were treated with EVs/siPD‐L1 or DP‐EVs/siPD‐L1. qPCR (Figure 4H) and western blot (Figure 4I,J) analyses confirmed a pronounced reduction in PD‐L1 mRNA and protein expression following DP‐EVs/siPD‐L1 treatment compared with EVs/siPD‐L1. Furthermore, incubation in mouse serum at 37°C for 24 h resulted in minimal siRNA leakage, as indicated by a largely unchanged EE compared with the untreated control (Figure S9A). Critically, serum‐incubated vesicles were still efficiently internalized by cells (Figure S9B) and retained high gene silencing activity (Figure S9C,D). Collectively, these results demonstrate that DP‐EVs function as a robust and stable nanocarrier platform for siRNA delivery, enabling efficient cellular internalization, enhanced endosomal escape, and effective gene silencing in both immune and tumor cells.
DP‐EVs/siPD‐L1 Activate Immune Response In Vivo
2.8
To assess the immunostimulatory effects of the engineered vesicles, we evaluated the immune response induced by DP‐EVs/siPD‐L1 complexes in healthy mice. Forty‐eight hours after subcutaneous injection, lymph nodes were harvested for flow cytometric analysis (Figures 5A and S12A). Mice treated with DP‐EVs/siPD‐L1 showed a marked increase in both the total DC (CD11c^+^) population and the proportion of mature DCs expressing high levels of CD80 and CD86, compared with those receiving EVs/siPD‐L1 (Figure 5B–D). Additionally, the in vivo knockdown efficiency was evaluated. Compared with the control groups (EVs/siNC and DP‐EVs/siNC), both EVs/siPD‐L1 and DP‐EVs/siPD‐L1 significantly downregulated PD‐L1 expression in lymph node tissues (Figure 5E). The simultaneous upregulation of DC maturation markers and suppression of PD‐L1 suggest that DP‐EVs/siPD‐L1 exert a synergistic immunostimulatory effect, facilitating both the activation of antigen‐presenting cells and the alleviation of immune checkpoint inhibition.
*DP‐EVs/siPD‐L1 can activate immune response in healthy mice. (A) The timeline process of the experimental scheme. (B–D) Detection of immune response in lymph nodes at 48 h after different treatment. (E) In vivo PD‐L1 knockdown efficiency detection in lymph nodes. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.001. N = 3.
In Vivo Antitumor Efficacy Evaluation of DP‐EVs/siPD‐L1
2.9
To assess the therapeutic potential of the engineered vesicles, an in vivo antitumor study was performed using E.G7‐OVA tumor‐bearing mice. DP‐EVs/siPD‐L1 complexes were administered subcutaneously following tumor inoculation (Figure 6A). Compared with control treatments, DP‐EVs/siPD‐L1 significantly suppressed tumor growth and prolonged overall survival. The results demonstrated that treatment with DP‐EVs/siPD‐L1 complexes led to a notable reduction in tumor growth and an increase in survival time. Remarkably, three out of seven mice achieved complete tumor regression, and the tumor growth inhibition (TGI) reached 75.8%, indicating potent therapeutic efficacy (Figure 6B–D). To further confirm the superiority of this delivery system, additional groups were included: free siPD‐L1, DP7‐C/siPD‐L1, and LNP/siPD‐L1 as a positive control. Among all formulations, DP‐EVs/siPD‐L1 produced the strongest antitumor response, comparable to that of the benchmark LNP/siPD‐L1, and significantly outperforming the other controls (Figure S10A–C). These findings highlight that DP‐EVs‐based nanoplatform provide a highly effective and biologically compatible platform for siRNA delivery, capable of achieving therapeutic efficacy equivalent to synthetic lipid nanoparticles.
*Antitumor efficacy of DP‐EVs/siPD‐L1 in E.G7‐OVA subcutaneous tumor mouse model. (A) The timeline process of the experimental scheme. (B) Individual tumor growth curves for E.G7‐OVA tumors in mice after different treatments (n = 7). (C) Average tumor growth curves for E.G7‐OVA tumors in mice after different treatments (n = 7). (D) Survival of mice with E.G7‐OVA tumors after various treatments (n = 7). (E and F) TUNEL, CD31, and Ki67 staining of E.G7‐OVA tumors after various treatments (n = 3). All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. Survival curves were obtained using the Kaplan–Meier method and compared by the log‐rank test. *p < 0.05, **p < 0.001.
Histological and systemic safety assessments further confirmed the biocompatibility of DP‐EVs/siPD‐L1 treatment. Hematoxylin and eosin (H&E) staining of major organs revealed no pathological abnormalities, and all groups showed stable increase of body weight during therapy, suggesting minimal systemic toxicity (Figure S11A,B). Immunohistochemical analyses provided mechanistic insights into the antitumor effect: CD31 staining indicated reduced microvessel density, Ki67 staining demonstrated decreased tumor cell proliferation, and TUNEL assays revealed enhanced apoptotic activity in tumors treated with DP‐EVs/siPD‐L1 (Figure 6E,F). Together, these results demonstrate that DP‐EVs/siPD‐L1 not only exert robust antitumor efficacy but also achieve it safely through the simultaneous inhibition of angiogenesis, suppression of tumor proliferation, and promotion of apoptosis.
Remodeling of the Immunosuppressive TME
2.10
To further elucidate how DP‐EVs/siPD‐L1 modulate the immune landscape, we analyzed immune cell populations in the tumor, lymph nodes, and spleen by flow cytometry (Figures 7A and S12B,C).
*Detection of TME reprogramming and immune response activation after different treatments. (A) The timeline process of the experimental scheme. (B) t‐Distributed stochastic neighbor embedding (t‐SNE) results of DCs, macrophages, T cells, and CD3− cells in E.G7‐OVA tumors after different treatments. (C) t‐SNE results of T cells in lymph nodes after different treatments. (D–K) Flow cytometry detection of DCs, macrophages, T cells, and CD3− cells in E.G7‐OVA tumors’ microenvironments after different treatments. (L) Schematic illustration of TME variation after DP‐EVs/siPD‐L1 administration. (M and N) Flow cytometry detection of T cells in lymph nodes after different treatments. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001. N = 3.
Within tumor tissues, we focused on T‐cell composition, dendritic‐cell maturation, macrophage polarization, and PD‐L1 expression dynamics. There was a significant knockdown of the PD‐L1 expression on nonimmune cells (CD3^−^) following the injection of siPD‐L1 delivered via EVs and DP‐EVs (Figure 7B,D). Correspondingly, the proportion of total DCs (CD11c^+^) and mature DCs (CD11c^+^CD80^+^CD86^+^) was markedly elevated after DP‐EVs/siPD‐L1 treatment, indicating enhanced antigen‐presenting capacity within the TME (Figure 7B,F). Additionally, a robust increase in cytotoxic T lymphocytes (CTLs; CD3^+^CD8^+^IFN‐γ^+^) (Figure 7B,G) and a significant decrease in regulatory T cells (Tregs; CD3^+^CD4^+^CD25^+^Foxp3^+^) (Figure 7B,H) were detected, reflecting a shift toward an effector‐dominant immune profile. DP‐EVs/siPD‐L1 treatment also altered macrophage polarization, elevating the M1 (CD11b^+^F4/80^+^CD86^+^) population while reducing M2 (CD11b^+^F4/80^+^CD206^+^) macrophages, resulting in a substantially higher M1/M2 ratio (Figure 7B,I–K). Collectively, these changes demonstrate that DP‐EVs/siPD‐L1 effectively reprogram the TME from immunosuppressive to immunostimulatory, characterized by PD‐L1 suppression, dendritic‐cell activation, T‐cell reinvigoration, and macrophage repolarization (Figure 7L).
To determine whether these effects extended to peripheral immune compartments, we next examined T‐cell subsets in lymph nodes and spleens. In lymph nodes, both EVs/siPD‐L1 and DP‐EVs/siPD‐L1 treatments markedly increased the proportion of CTLs while significantly reducing the frequency of Tregs (Figure 7C,M,N). A similar trend was observed in the spleen, where CTL levels were substantially elevated following treatment (Figure S13A,B). Notably, a decrease in Treg frequency was observed only in the DP‐EVs/siPD‐L1 group, whereas Treg levels in other treatment groups remained largely unchanged (Figures S12A and S13C). These results suggest that DP‐EVs/siPD‐L1 not only remodel the local TME but also promote systemic immune activation, thereby potentiating coordinated antitumor immunity.
Altered miRNA Cargo in DP‐EVs Contributes to Their Enhanced Antitumor Activity
2.11
To elucidate the molecular basis underlying the superior antitumor efficacy of DP‐EVs, we examined the miRNA profiles identified by sRNA sequencing (Figure S14A). qPCR validation confirmed a subset of miRNAs that were differentially expressed in DP‐EVs compared with native EVs, including miR‐128‐2‐5p, miR‐142a‐5p, miR‐181b‐1‐3p, miR‐182‐5p, miR‐211‐5p, miR‐328‐3p, and miR‐451a (Figure S14B). These results indicate that DP7‐C treatment during EV biogenesis substantially alters the RNA cargo composition, potentially endowing DP‐EVs with unique biological functions. Among these, miR‐451a has been widely recognized as a tumor suppressor in multiple malignancies [38, 39, 40]. To verify its functional relevance, miR‐451a mimics were transfected into E.G7‐OVA cells, and the expression of epithelial–mesenchymal transition (EMT) markers was subsequently analyzed. Consistent with the effects observed in DP‐EV treated cells, miR‐451a overexpression increased E‐cadherin levels while reducing N‐cadherin and vimentin expression (Figure S14C), suggesting that miR‐451a enrichment in DP‐EVs contributes to the inhibition of EMT and the attenuation of tumor invasiveness. Besides, miR‐142a‐5p, which has been implicated in promoting M2‐like macrophage polarization [41], showed a distinct regulatory trend. Treatment of macrophages with synthetic miR‐142a‐5p elevated the proportion of M2‐type macrophages, whereas DP‐EVs, compared with EVs, significantly reduced M2‐like polarization (Figure S14D). These results imply that the downregulation of miR‐142a‐5p within DP‐EVs may partially account for their ability to repolarize macrophages toward an M1 phenotype. Collectively, these findings demonstrate that DP7‐C induced remodeling of EV miRNA cargo, characterized by the enrichment of tumor‐suppressive miRNAs and depletion of protumorigenic species, plays a pivotal role in the enhanced antitumor activity of DP‐EVs.
Discussion
3
In this study, we developed a novel class of engineered EVs (DP‐EVs) by culturing tumor cells with the immunomodulatory peptide DP7‐C. DP‐EVs preserved the characteristic morphology and surface charge of native EVs but exhibited a slightly larger diameter and distinct molecular cargos, indicating that DP7‐C reshapes EV biogenesis and vesicle composition. Functionally, DP‐EVs showed markedly enhanced cellular uptake via macropinocytosis, caveolin and clathrin‐mediated pathways, as well as superior endosomal escape capability. These improvements enabled DP‐EVs to serve as efficient vehicles for siPD‐L1 delivery while maintaining intrinsic immunostimulatory properties such as DC maturation and macrophage M1 polarization. Consequently, siPD‐L1‐loaded DP‐EVs (DP‐EVs/siPD‐L1) achieved potent tumor suppression through a dual mechanism involving PD‐L1 blockade and immune activation‐driven remodeling of the TME.
The present work presents several important mechanistic and conceptual advances. First, a simple incubation of tumor cells with DP7‐C was sufficient to endow secreted vesicles with enhanced targeting toward both lymph nodes and tumor tissues, accompanied by improved uptake efficiency by donor tumor cells. This observation aligns with previous findings that cellular stimulation or microenvironmental modulation can generate exosomes with distinct functional phenotypes [42]. To elucidate the molecular basis underlying these enhanced biological functions, we investigated how DP7‐C treatment influences EV properties. DP7‐C is a cationic amphipathic peptide capable of associating with negatively charged phospholipids and glycoproteins on the plasma membrane. During EV biogenesis, DP7‐C partially incorporates into the cell membrane and is subsequently integrated into the forming intraluminal vesicles. This process likely alters the lipid microdomain organization, increases surface fluidity, and slightly raises the surface charge, as reflected by the higher zeta potential and larger particle size of DP‐EVs (Figure 1D,E). Such membrane remodeling enhances electrostatic and hydrophobic interactions between DP‐EVs and recipient cell membranes, facilitating stronger vesicle–cell adhesion and internalization. In parallel, proteomic and miRNA analyses revealed that DP7‐C treatment enriched pathways related to actin cytoskeleton regulation, Fc gamma R‐mediated phagocytosis, and endocytosis, signaling processes known to promote receptor‐mediated uptake and intracellular trafficking. These combined effects provide a molecular explanation for the observed dual targeting of DP‐EVs to both dendritic and tumor cells.
In addition to altered surface properties, DP‐EVs display a broadened internalization profile. Unlike conventional EVs, which primarily rely on macropinocytosis and caveolin‐mediated entry [43], DP‐EVs also utilize a clathrin‐dependent pathway, a feature likely attributed to membrane remodeling by DP7‐C, as our previous work demonstrated that DP7‐C/peptide and DP7‐C/siRNA complexes were internalized by DCs via both caveolin and clathrin‐dependent routes [28, 37]. This broadened uptake mechanism, together with their superior endosomal escape capability, ensures more efficient cytoplasmic release of therapeutic cargos such as siRNA [44]. These properties represent a substantial improvement over natural EVs and may significantly enhance the efficacy of EV‐based drug delivery systems.
Importantly, DP‐EVs/siPD‐L1 show a remarkable efficacy for tumor immunotherapy. Two major factors contribute to the superior antitumor efficacy. First, their dual immunomodulatory‐delivery functionality integrates immune activation with efficient siRNA transport. The intrinsic presence of TAAs enables DP‐EVs to activate antigen‐presenting cells, a feature absent in synthetic carriers like liposomes or lipid nanoparticles [45]. Unlike these carriers, which are primarily designed for nucleic acid delivery and lack intrinsic immunostimulatory capacity, DP‐EVs possess a dual functionality. As immune activators, DP‐EVs promote DC maturation and M1‐type macrophage polarization in vitro; as delivery vectors, they efficiently transport siPD‐L1 to target cells, leading to potent PD‐L1 silencing and enhanced cytotoxic T‐cell responses [46]. This synergy establishes a self‐amplifying “blockade‐and‐activation” cycle that drives robust immune remodeling and tumor regression in vivo. Second, DP‐EVs exhibit selective miRNA remodeling. A cluster analysis of the differentially expressed miRNAs is presented in Figure S14E. Notably, the majority of downregulated species are frequently linked to the promotion of tumor progression [41, 47, 48, 49, 50, 51, 52, 53, 54, 55], whereas the upregulated miRNAs in DP‐EVs (e.g., mmu‐miR‐211‐5p and mmu‐miR‐328‐3p) are associated with tumor‐suppressive functions [56, 57, 58, 59, 60, 61, 62, 63, 64, 65]. This selective cargo reprogramming likely contributes to the enhanced immunomodulatory and antitumor properties of DP‐EVs, providing a second mechanistic layer beyond surface modulation.
While this study demonstrates the therapeutic potential of DP‐EVs, several limitations should be acknowledged, which also present opportunities for future research. First, the precise biological functions of the differentially expressed proteins and miRNAs identified in our omics analysis remain to be fully elucidated. While their association with enhanced immune activation is evident, direct functional validation is necessary to establish causal relationships. Second, the molecular mechanisms underlying DP‐EV mediated DC maturation and immune activation, particularly the specific signaling pathways involved, require more detailed dissection. Although our data suggest involvement of pathways like Fc gamma R‐mediated phagocytosis, further mechanistic studies are needed to confirm these pathways’ roles. Finally, and most critically, the scalable production and standardization of engineered EVs remain a significant challenge for clinical translation. Current manufacturing methods may not be sufficient for large‐scale, reproducible production of clinical‐grade DP‐EVs, which could impact batch‐to‐batch consistency and therapeutic efficacy.
In conclusion, this work establishes DP‐EV/siPD‐L1 as a novel therapeutic vaccine, a next‐generation EV‐based platform that synergistically unites efficient siRNA delivery with potent immune activation. By simultaneously targeting the PD‐L1 checkpoint to alleviate immunosuppression and reprogramming the TME through active DC engagement, this strategy demonstrates a dual functionality, acting both prophylactically and therapeutically against tumors. The engineered DP‐EV core not only serves as a targeted delivery vehicle but also functions as an intrinsic immune adjuvant, thereby enhancing the specificity and potency of the antitumor response. Future studies optimizing their production, cargo design, and mechanism of action will pave the way for their advancement toward precision immunotherapy and improved patient outcomes.
Materials and Methods
4
Cell Lines and Animal Models
4.1
The E.G7‐OVA and 4T1 cell line was obtained from the American Type Culture Collection. BMDCs and BMDMs were generated from female C57BL/6J female mice (4–6 weeks old; Vital River Laboratory Animal Technology Co., Ltd., Beijing, China). All cells were cultured at 37°C in 5% CO_2_ with Roswell Park Memorial Institute (RPMI)‐1640 medium containing 10% fetal bovine serum (FBS), 100 units/mL streptomycin and 100 units/mL penicillin (PS). All animal experiments were performed in accordance with protocols approved by the Ethics Committee of Sichuan University, following institutional and national guidelines for the care and use of laboratory animals.
BMDCs and BMDMs were prepared as previously described [28]. Briefly, bone marrow cells were flushed from the femurs and tibias of C57BL/6J mice and treated with red blood cell lysis buffer. The remaining cells were cultured in complete RPMI‐1640 medium supplemented with either 20 ng/mL granulocyte–macrophage colony‐stimulating factor (GM‐CSF; PrimeGene Biotechnology, Shanghai, China) and 20 ng/mL recombinant mouse IL‐4 to induce BMDC differentiation, or with 20 ng/mL macrophage CSF to generate BMDMs. The culture medium was refreshed every 2 days, and cells were used for experiments after 7 days of differentiation.
Cell Viability Assessment (CCK‐8 Assay)
4.2
E.G7‐OVA cells were seeded in 96‐well plates at a density of 1 × 10^5^ cells per well and allowed to adhere for 24 h. The cells were then treated with various concentrations of DP7‐C for an additional 24 h. After treatment, 10 µL of Cell Counting Kit‐8 (CCK‐8) reagent was added to each well, followed by incubation at 37°C for 30 min. Cell viability was quantified by measuring the absorbance at 450 nm using a microplate reader (Tecan Infinite 200 Pro, M200 PRO/F200 PRO, Austria).
Preparation of EVs and DP‐EVs
4.3
Engineered EVs (DP‐EVs) were isolated from E.G7‐OVA cells cultured in serum‐free medium containing either 0 or 50 µg/mL DP7‐C. After 24 h of incubation, the conditioned medium was collected and processed by differential ultracentrifugation as previously described [66]. Briefly, the supernatant was sequentially centrifuged at 300×g for 10 min, 2000×g for 10 min, and 10,000×g for 30 min, to remove cells and debris. The clarified medium was then passed through a 0.22 µm filter (Merck Millipore, Germany) and ultracentrifuged at 100,000×g for 120 min (Beckman Coulter Optima XPN; SW32Ti rotor; USA). The resulting pellet was resuspended in PBS, followed by a second ultracentrifugation at 100,000×g for 120 min to eliminate residual contaminants. Purified EVs or DP‐EVs were finally resuspended in PBS, aliquoted, and stored at −80°C to avoid repeated freeze–thaw cycles.
Characterization of EVs and DP‐EVs
4.4
The morphology of EVs and DP‐EVs was examined using TEM (JSM‐7500F; JEOL, Japan). The particle size distribution and zeta potential were measured by NTA (ZetaView; Particle Metrix; Germany). For western blotting, EVs and DP‐EVs were lysed in RIPA buffer (Beyotime, China), and the protein concentration was quantified using a BCA Protein Assay Kit (Beyotime). Equal amounts of protein were separated by 10% SDS‐PAGE and transferred onto polyvinylidene fluoride (PVDF) membranes. After blocking with 5% nonfat milk, membranes were incubated with primary antibodies against CD9, CD63, TSG101, and GAPDH (HuaBio, China), followed by HRP‐conjugated secondary antibodies (HuaBio). Protein bands were visualized using enhanced chemiluminescence (ECL; Thermo Fisher Scientific, USA) and imaged with a western blot detection system (e‐BLOT Life Science, Shanghai, China).
To compare molecular cargos, small RNA sequencing and proteomic analysis of EVs and DP‐EVs were performed by Novogene Co., Ltd. (Beijing, China). For small RNA sequencing, total RNA was extracted using an exosomal RNA extraction kit (EZB, China) and sequenced on the Illumina SE50 platform. For proteomic profiling, purified EVs and DP‐EVs were analyzed by label‐free quantitative proteomics, and the resulting data were processed using NovaMagic Plus (novogene.com).
In Vivo Biological Distribution
4.5
To evaluate the biodistribution of EVs, both EVs and DP‐EVs were labeled with the near‐infrared fluorescent dye DiR (Yeasen, China) at 37°C for 30 min. Unbound dye was removed by ultrafiltration, and the labeled vesicles were resuspended in PBS for subsequent use. Fifty micrograms of DiR‐labeled EVs or DP‐EVs were subcutaneously injected at the base of the tail in E.G7‐OVA tumor‐bearing mice (n = 3). After 24 h, the mice were sacrificed, and major organs were excised and imaged using an IVIS Lumina III imaging system (PerkinElmer, USA) to assess fluorescence distribution. Tumor tissues and draining lymph nodes were immediately harvested, embedded in OCT compound, and snap‐frozen in liquid nitrogen. Frozen sections (8 µm) were prepared using a cryostat microtome (Leica CM1950, Germany), fixed with 4% paraformaldehyde, stained with DAPI (Beyotime), and visualized under a confocal laser scanning microscope (LSM 900; Zeiss, Germany). To further investigate the cellular targeting mechanism, lymph nodes and tumor tissues were digested into single‐cell suspensions and stained with antibodies against CD45, CD11c, and CD103 for flow cytometric analysis using a NovoCyte Advanteon Dx flow cytometer (Agilent, USA).
In Vitro Uptake Efficiency
4.6
BMDCs, BMDMs, and E.G7‐OVA cells were seeded in 24‐well plates at a density of 1 × 10^5^ cells per well and cultured for 24 h. The cells were then incubated with 1 µg of DiO‐labeled EVs or DP‐EVs (Yeasen) for an additional 24 h. After incubation, cellular uptake was visualized using a fluorescence microscope (Nikon Eclipse Ti, Japan). The cells were subsequently collected, washed with PBS, and analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent) to quantify uptake efficiency.
Uptake Pathway Analysis
4.7
To investigate the mechanisms of EV and DP‐EV uptake by DCs, BMDCs (1 × 10^5^ cells per well) were seeded into 24‐well plates and cultured for 24 h. Cells were then pretreated for 2 h with specific endocytic inhibitors, including amiloride (20 µM; macropinocytosis inhibitor), chlorpromazine (5 µM; clathrin‐mediated endocytosis inhibitor), and genistein (30 µM; caveolin‐mediated endocytosis inhibitor) (all from Selleck, USA). After pretreatment, the inhibitor‐containing medium was replaced with fresh medium containing 1 µg of DiO‐labeled EVs or DP‐EVs, followed by incubation for an additional 24 h. The cells were then collected, washed with PBS, and analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent) to quantify uptake efficiency.
For colocalization analysis, BMDCs (1 × 10^5^ cells per well) were cultured on sterilized glass slides in 24‐well plates for 24 h, followed by incubation with 1 µg of DiO‐labeled EVs or DP‐EVs for 4 h. To visualize the endocytic pathways, cells were stained with specific fluorescent probes: Dextran Texas Red (20 mg/mL, 37°C for 20 min; Invitrogen, USA) for macropinocytosis, CT‐B (10 µg/mL, 4°C for 15 min; Invitrogen) for caveolin‐mediated uptake, and Transferrin‐Alexa Fluor 555 (50 µg/mL, 37°C for 2 h; Jackson ImmunoResearch, USA) for clathrin‐mediated uptake. After staining, cells were fixed with 4% paraformaldehyde, mounted with an anti‐fade medium containing DAPI (Solarbio, China), and imaged using a confocal laser scanning microscope (LSM 900; Zeiss).
Endosomal and Lysosomal Escape Assay
4.8
To investigate the endosomal and lysosomal escape of EVs after cellular uptake, BMDCs (1 × 10^5^ cells per well) were seeded onto glass coverslips in 24‐well plates and cultured for 24 h. The cells were then incubated with 1 µg of DiR‐labeled EVs or DP‐EVs for endocytosis tracking. For lysosomal staining, LysoTracker Green (Beyotime) was added to the wells at 4 or 22 h postincubation, followed by an additional 2 h incubation at 37°C. After staining, the cells were washed, fixed with 4% paraformaldehyde, counterstained with DAPI, and mounted for confocal imaging. For endosome staining, cells were fixed with 4% paraformaldehyde and permeabilized using 0.5% Triton X‐100 for 20 min at room temperature. After blocking with 5% BSA for 20 min, cells were incubated with the primary antibody against EEA1 (1:100; Invitrogen) for early endosomes or FITC‐labeled LAMP1 (1:100; BD Biosciences, USA) for late endosomes. For EEA1 staining, samples were subsequently incubated with an FITC‐conjugated secondary antibody for 40 min at room temperature. Finally, the cells were counterstained with DAPI and imaged using a confocal laser scanning microscope (LSM 900; Zeiss).
In Vitro Differentiation and Maturation of BMDCs
4.9
To evaluate the effects of EVs and DP‐EVs on the differentiation and maturation of BMDCs, cells were seeded into 24‐well plates at a density of 5 × 10^5^ cells/mL and incubated with 1 µg of EVs or DP‐EVs for 24 h. After incubation, the cells were collected, washed twice with cold PBS containing 1% FBS, and stained with fluorophore‐conjugated antibodies against CD11c, CD80, and CD86 (BD Biosciences) at 4°C for 40 min in the dark. The cells were then washed and analyzed using a flow cytometer (NovoCyte Advanteon Dx; Agilent) to determine the proportion of mature BMDCs.
In Vitro Migration and Mechanistic Study of BMDCs
4.10
To assess the effects of EVs and DP‐EVs on BMDC migration, cells were seeded into 24‐well plates at a density of 5 × 10^5^ cells/mL and treated with 1 µg of EVs or DP‐EVs for 24 h. After incubation, BMDCs were harvested and transferred to Transwell inserts (Corning, USA). The upper chamber contained 5 × 10^5^ BMDCs in 100 µL serum‐ and antibiotic‐free RPMI‐1640 medium, while the lower chamber was filled with 100 µL of RPMI‐1640 supplemented with 10% FBS, 1% PS–streptomycin, 250 ng/mL CCL19, and 250 ng/mL CCL21 (Sino Biological, China). After 24 h, cells that migrated to the lower chamber and those remaining in the upper chamber were collected and counted using an automated cell counter (Countstar IM1200; Shanghai, China). To explore the molecular mechanisms underlying EV‐induced BMDC migration, CCR7 expression was analyzed by quantitative real‐time PCR (qRT‐PCR), western blotting, and immunofluorescence staining.
For qRT‐PCR, BMDCs were seeded in 12‐well plates (5 × 10^5^ cells/mL) and treated with 2 µg of EVs or DP‐EVs for 24 h. Total RNA was extracted using an Animal Total RNA Isolation Kit (Foregene, China), and complementary DNA (cDNA) was synthesized with HiScript III RT SuperMix for qPCR (Vazyme, China). qPCR was performed using ChamQ Universal SYBR qPCR Master Mix (Vazyme). Primer sequences were listed in Table S1.
For western blotting, BMDCs were seeded in six‐well plates (5 × 10^5^ cells/mL) and treated with 4 µg of EVs or DP‐EVs for 24 h. Total proteins were extracted, separated by SDS‐PAGE (FastPAGE Precast Gel; Tsingke Biotechnology, China), and transferred to PVDF membranes. Membranes were incubated with primary antibodies against CCR7, AKT, p‐AKT, PI3K, p‐PI3K, JNK, p‐JNK, MAPK, p‐MAPK, ERK, p‐ERK, p65, and p‐p65 (HuaBio; Cell Signaling Technology [CST], USA), followed by HRP‐conjugated secondary antibodies (Abcam, USA). Protein bands were visualized using an ECL detection system (Millipore, USA).
For immunofluorescence staining, 1 × 10^5^ BMDCs were cultured on glass slides in 24‐well plates and incubated with 1 µg of DiR‐labeled EVs or DP‐EVs for 24 h. Cells were then fixed with 4% paraformaldehyde, stained with anti‐CCR7 (HuaBio) and an iFluor 488–conjugated goat anti‐rabbit IgG secondary antibody (HuaBio) at 37°C for 40 min, counterstained with DAPI (Beyotime), and imaged using a confocal laser scanning microscope (LSM 900; Zeiss).
For flow cytometry, 1 × 10^5^ BMDCs were cultured in 24‐well plates and incubated with 1 µg EVs or DP‐EVs for 24 h. Cells were then harvested and stained with fluorophore‐conjugated antibodies against CCR7 (BD Biosciences) at 4°C for 40 min in the dark. The stained cells were washed, resuspended in PBS, and analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent).
In Vitro Polarization of BMDMs
4.11
To evaluate the effects of EVs and DP‐EVs on macrophage polarization, BMDMs were seeded into 24‐well plates at a density of 1 × 10^6^ cells/mL and treated with 1 µg of EVs or DP‐EVs for 24 h. After incubation, the cells were collected, washed twice with cold PBS containing 1% FBS, and stained with fluorophore‐conjugated antibodies against CD11b, F4/80, CD86 (BD Biosciences), and CD206 (BioLegend, USA) at 4°C for 40 min in the dark. The stained cells were washed, resuspended in PBS, and analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent) to determine the ratio of M1‐like (CD11b^+^F4/80^+^CD86^+^) to M2‐like (CD11b^+^F4/80^+^CD206^+^) macrophages. To further analyze the molecular characteristics of macrophage polarization, qRT‐PCR was performed to assess the expression of polarization‐related genes. The primer sequences were listed in Table S1 and the qRT‐PCR procedure followed the same protocol described in Section 4.10.
Loading of siRNA Into EVs
4.12
siRNA was loaded into EVs or DP‐EVs using an electroporation method. Briefly, siRNA and EVs were mixed in Bio‐Rad electroporation buffer at a mass ratio of 1:4 and kept on ice for 1 h. Electroporation was performed using a Gene Pulser Xcell Electroporation System (Bio‐Rad, USA) at 200 V, ∞ Ω, and 125 µF (CE mode). The final EV concentration was maintained at 0.5 µg/µL. After electroporation, the mixture was incubated on ice for 30 min to allow membrane recovery, followed by ultracentrifugation at 100,000×g for 120 min to remove unencapsulated siRNA. The resulting EV/siRNA and DP‐EV/siRNA complexes were resuspended in PBS for in vitro and in vivo experiments.
LNPs encapsulating siRNA were prepared using a microfluidic mixing method. Briefly, the aqueous phase was composed of siRNA dissolved in 10 mM citrate buffer (pH 4.0). The organic phase consisted of lipids (SM‐102, DSPC, cholesterol, PEG‐lipid) dissolved in absolute ethanol at a molar ratio of 50:10:38.5:1.5. Both phases were loaded into a microfluidic mixing device (Micro&Nano, China) and rapidly mixed at a volumetric flow ratio of aqueous:organic = 3:1, with a total flow rate of 12 mL/min. The lipid‐to‐siRNA ratio was set to a mass ratio of 15:1. Immediately after mixing, the resulting LNP dispersions were collected and subjected to buffer exchange via ultrafiltration using Amicon Ultra centrifugal filters (100 kDa MWCO) to remove ethanol and adjust the pH.
Characterization of EV/siRNA and DP‐EV/siRNA
4.13
The siRNA EE was determined using an ultracentrifugation‐based method. Briefly, EVs/cy5‐siRNA and DP‐EVs/cy5‐siRNA complexes were resuspended in 8.5 mL PBS and centrifuged at 100,000×g for 70 min. The pellet was resuspended, and cy5 fluorescence was quantified using a microplate reader (excitation/emission: 650/670 nm). A standard curve was generated using serially diluted cy5‐siRNA (6.25–0 pg/µL). EE (%) = (encapsulated siRNA/total siRNA) × 100%.
To assess nuclease protection, free siRNA (1 µg), EV/siRNA (1 µg), and DP‐EV/siRNA (1 µg) were incubated with RNase A (0.1 µg) at 37°C for 0, 0.5, 1, 2, or 4 h. Reactions were terminated by adding SDS (final 0.5%) and incubating for 10 min. Samples were analyzed by agarose gel electrophoresis (120 V, 10 min) and visualized using a ChemiDoc XRS Imaging System (Bio‐Rad).
For serum stability evaluation, EV/cy5‐siRNA and DP‐EV/cy5‐siRNA were incubated in 10% mouse serum at 37°C for 24 h. The serum‐treated complexes were subsequently applied to BMDCs and E.G7‐OVA cells to assess siRNA leakage and cellular uptake efficiency by flow cytometry. Similarly, EV/siPD‐L1 and DP‐EV/siPD‐L1 complexes were used to evaluate retained gene silencing activity under serum exposure.
In Vitro Gene Silencing Efficiency of EVs/siPD‐L1 by qRT‐PCR
4.14
To determine PD‐L1 gene silencing efficiency, BMDCs (5 × 10^5^ cells/mL) and E.G7‐OVA cells (2 × 10^5^ cells/mL) were seeded in 12‐well plates and treated with PBS, EV (4 µg)/siNC (1 µg), EV (4 µg)/siPD‐L1 (1 µg), DP‐EV (4 µg)/siNC (1 µg), or DP‐EV (4 µg)/siPD‐L1 (1 µg) for 24 h. The sequences of the siPD‐L1 oligonucleotides were: sense: 5′‐GUCCUCCUCUUCUUGAGAATT‐3′; antisense: 5′‐UUCUCAAGAAGAGGAGGACTT‐3′ (Ketai, Shanghai). Total RNA was extracted to perform qRT‐PCR, followed the same protocol described in Section 4.10 and the primers sequences were listed in Table S1.
In Vitro Gene Silencing Efficiency of EVs/siPD‐L1 by Western Blot Analysis
4.15
To evaluate PD‐L1 protein silencing, BMDCs (5 × 10^5^ cells/mL) and E.G7‐OVA cells (2 × 10^5^ cells/mL) were seeded into six‐well plates and treated with PBS, EV (8 µg)/siNC (2 µg), EV (8 µg)/siPD‐L1 (2 µg), DP‐EV (8 µg)/siNC (2 µg), or DP‐EV (8 µg)/siPD‐L1 (2 µg) for 48 h. Western blotting was performed following the procedure described in Section 4.4. Newly applied antibodies included anti‐PD‐L1 (CST) and anti‐GAPDH (HuaBio).
Detection of EVs and DP‐EVs‐Activated Immune Responses
4.16
To evaluate immune activation following EV administration, female C57BL/6J mice (6–8 weeks old) were subcutaneously injected at the tail base with PBS, EV/siNC, EV/siPD‐L1, DP‐EV/siNC, or DP‐EV/siPD‐L1 formulations (10 µg EVs per mouse in 100 µL PBS). After 48 h, the mice were euthanized, and lymph nodes were harvested and mechanically dissociated into single‐cell suspensions. The cells were then stained with fixable viability stain (FVS) 780, anti‐mouse CD45, anti‐mouse CD11c, anti‐mouse CD80, anti‐mouse CD86 (BD Biosciences), and anti‐mouse PD‐L1 (BioLegend) at 4°C for 40 min in the dark. After washing and fixation, samples were analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent) to determine the activation status of DCs and the expression of PD‐L1.
In Vivo Antitumor Efficacy
4.17
To evaluate the antitumor efficacy of siPD‐L1‐loaded EVs, female C57BL/6J mice (6–8 weeks old) were subcutaneously inoculated with 7 × 10^5^ E.G7‐OVA tumor cells on Day 0. On Days 4, 11, and 18, mice received subcutaneous injections of PBS, EV/siNC, EV/siPD‐L1, DP‐EV/siNC, or DP‐EV/siPD‐L1 formulations (10 µg EVs per mouse in 100 µL PBS) at the base of the tail. Each group contained seven mice (n = 7). Tumor growth, body weight, and survival were monitored every 2 days. Tumor volume (TV) was calculated using the formula:
and TGI was determined as:
Mice were euthanized when TVs exceeded 1500 mm^3^ in accordance with ethical guidelines to minimize suffering.
Histological and Apoptosis Analysis
4.18
For histological evaluation, female C57BL/6J mice (6–8 weeks old) were subcutaneously inoculated with 7 × 10^5^ E.G7‐OVA tumor cells on Day 0. Mice were subsequently administered PBS, EV/siNC, EV/siPD‐L1, DP‐EV/siNC, or DP‐EV/siPD‐L1 (10 µg EVs per mouse in 100 µL PBS; n = 3) via subcutaneous injection at the tail base on Days 4, 11, and 18. On Day 20, mice were euthanized, and the half of tumor tissues collected on Day 20 were fixed in 4% paraformaldehyde, embedded in paraffin, sectioned, and stained for CD31 and Ki67 (Abcam, UK) to assess angiogenesis and tumor proliferation, respectively. Briefly, tissue sections were incubated overnight with primary antibodies at 4°C, followed by HRP‐conjugated goat anti‐rabbit IgG at 37°C for 1 h. Visualization was achieved with DAB substrate, and slides were mounted with neutral resin. Images were acquired using a slide scanner (Pannoramic MIDI; 3DHISTECH Ltd., Hungary). Apoptotic cells were detected using a TUNEL Apoptosis Detection Kit (Vazyme) according to the manufacturer's protocol, and fluorescence images were captured using a confocal laser scanning microscope (LSM900; Zeiss).
Profiling of Immune Response and TME
4.19
For immune activation and TME remodeling evaluation, lymph nodes, spleens, and the other half of tumor tissues were harvested and dissociated into single‐cell suspensions for flow cytometry analysis (n = 3). Two antibody panels were employed to characterize adaptive and innate immune responses: T cell activation panel: FVS 780, anti‐mouse CD45, CD3, CD4, CD8, CD25, Foxp3, IFN‐γ, and PD‐L1 (BD Biosciences). These markers were used to quantify CTLs, Tregs, and to confirm in vivo PD‐L1 knockdown efficiency. Innate immune panel: FVS 780, anti‐mouse CD11b, CD11c, CD86, and CD206 (BD Biosciences), used to assess DC maturation and macrophage polarization within the TME. After staining at 4°C for 40 min in the dark, samples were fixed and analyzed by flow cytometry (NovoCyte Advanteon Dx; Agilent). The resulting data were processed to evaluate immune cell activation profiles across tumor, lymph node, and spleen compartments.
In Vivo Safety Evaluation
4.20
To assess systemic safety, female C57BL/6J mice (6–8 weeks old) were subcutaneously injected with PBS, EV/siNC, EV/siPD‐L1, DP‐EV/siNC, or DP‐EV/siPD‐L1 (10 µg EVs per mouse in 100 µL PBS) on Days 0, 7, and 14, simulating the therapeutic dosing schedule. On Day 16, major organs—including the heart, liver, spleen, lung, and kidney—were collected, fixed in 4% paraformaldehyde, embedded in paraffin, and stained with H&E following the manufacturer's protocol (Solarbio). Histological sections were imaged using a slide scanning system (Pannoramic MIDI; 3DHISTECH Ltd., Hungary) to evaluate tissue morphology and potential treatment‐related toxicity.
qPCR Validation of Differentially Expressed miRNA in EVs and DP‐EVs
4.21
To validate the differential expression of miRNAs identified by small RNA sequencing, qPCR was performed. Briefly, total small RNAs were extracted from both EVs and DP‐EVs, and cDNA was synthesized using the miRNA first Strand cDNA Synthesis Kit (by stem‐loop) (Vazyme). qPCR was subsequently conducted using the miRNA Universal SYBR qPCR Master Mix (Vazyme) according to the manufacturer's protocol. Primer sequences used for amplification are listed in Tables S2 and S3.
Evaluation of the Antitumor Effect of miR‐451a in E.G7‐OVA Cells
4.22
To evaluate the antitumor role of miR‐451a, which was found to be upregulated in DP‐EVs, E.G7‐OVA cells were treated with EVs (5 µg), DP‐EVs (5 µg), Lipo2000/mimic NC (4 µg/2 µg), or Lipo2000/miR‐451a mimic (4 µg/2 µg). After 48 h of incubation, total cellular protein was extracted, and EMT markers, including E‐cadherin, N‐cadherin, and vimentin, were analyzed by western blotting using specific antibodies (CST). The sequences of the miR‐451a mimic used in this study were: sense, 5′‐AAACCGUUACCAUUACUGAGUU‐3′; antisense, 5′‐CUCAGUAAUGGUAACGGUUUUU‐3′ (GenePharma, Shanghai, China).
Macrophage Polarization Verification of miR‐142a‐5p
4.23
To confirm the role of miR‐142a‐5p in macrophage polarization, BMDMs were treated with EVs (5 µg), DP‐EVs (5 µg), Lipo2000/mimic NC (4 µg/2 µg), or Lipo2000/miR‐142a‐5p mimic (4 µg/2 µg). After 24 h of incubation, cells were collected for flow cytometry analysis to quantify M2‐like macrophages using fluorophore‐conjugated antibodies against CD11b, F4/80, and CD206 (BD Biosciences). In parallel, total RNA was extracted from each group to determine CD206 expression by qPCR, and the primer sequences were listed in Table S1. The sequences of the miR‐142a‐5p mimic were: sense, 5′‐CAUAAAGUAGAAAGCACUACU‐3′; antisense, 5′‐UAGUGCUUUCUACUUUAUGUU‐3′ (GenePharma).
Statistical Analysis
4.24
All statistical analyses were performed using GraphPad Prism 8.0 (GraphPad Software, USA). Data are presented as mean ± standard deviation (s.d.) unless otherwise specified. Comparisons among multiple groups were conducted using one‐way analysis of variance (ANOVA) followed by Tukey's posthoc multiple comparison test, while comparisons between two groups were analyzed using Student's t‐test. Survival curves were generated using the Kaplan–Meier method and analyzed with the log‐rank test. A p value of <0.05 was considered statistically significant. Significance levels were denoted as follows: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
Author Contributions
Li Yang, Rui Zhang, and Yusi Wang designed the study. Yusi Wang and Rui Zhang responsible for all experiments and articles. Xuejing Zhou, Lin Tang, and Die Hu helped Yusi Wang to perform the tumor model. Yuling Yang and Yibing Zhang helped Yusi Wang to perform the in vitro experiment. Bailing Zhou contributed to manuscript corrections. All authors read and approved the final manuscript.
Funding Information
This work was supported by the National Key Research and Development Program of China (No. 2023YFC3405200), the National Natural Science Foundation of China (No. 82471854), the 1.3.5 project for disciplines of excellence, West China Hospital, Sichuan University (No. ZYGD23008), Sichuan Science and Technology Program (2023YFS0001), National Clinical Research Center for Geriatrics, West China Hospital, Sichuan University (Z2023LC001), and The Frontiers Medical Center, Tianfu Jincheng Laboratory Foundation (TFJC2023010005).
Conflicts of Interest
The authors declare that they have no competing interests.
Ethics Statement
All animal experiments in this study were approved by the Institutional Animal Care and Use Committee of Sichuan University (ethical approval number: 20230307056).
Supporting information
Table S1: Primer sequences used for qRT‐PCR analysis of target mRNAs. Table S2: Primer sequences used for cDNA synthesis of target microRNAs and U6. Table S3: Primer sequences used for qRT‐PCR of target microRNAs and U6. Figure S1: In vivo distribution of EVs and DP‐EVs in E.G7‐OVA bearing mice. (A) Cell viability of E.G7‐OVA after 24 h incubation with different DP7‐C concentration. (B) In vivo biodistribution after subcutaneous injection of DIR‐EVs and DIR‐DP‐EVs (n = 3). Figure S2: DP‐EVs exhibited higher accumulation in tumor and lymph nodes in 4T1 model. (A and B) In vivo biodistribution after 24 h of subcutaneous injection of DIR‐EVs and DIR‐DP‐EVs in 4T1 baring mice (n = 3). All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05. Figure S3: Endosomal escape of EV and DP‐EV in DCs. (A) Representative colocalization images after incubation in BMDCs of early endosomes, late endosomes, and lysosomes with EVs and DP‐EVs at 6 h. (B) Representative colocalization images of early endosomes, late endosomes, and lysosomes after incubation in BMDCs with EVs and DP‐EVs at 24h. Scale bars = 10 µm. Figure S4: CCR7 expression in dendritic cells (DCs) treated with EVs or DP‐EVs. (A) Flow cytometry analysis of CCR7 expression on CD11c^+^ DCs after treatment with EVs or DP‐EVs. (B) Immunofluorescence staining confirming CCR7 protein expression. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Scale bars = 20 µm. Figure S5: DP‐EVs promote macrophage M1 polarization. (A) Flow cytometric analysis of surface markers CD80 and CD86 on macrophages treated with PBS, EV, and DP‐EV. Representative density plots show the expression levels. (B) Quantification of the percentages of CD206^+^ and CD86^+^ cells in CD11b^+^ F480^+^ cells. (C) mRNA expression levels of M1 macrophage‐associated genes by qRT‐PCR in macrophages treated as indicated. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Figure S6: Characterization and endosomal escape of EV/siRNA. (A) TEM images EV and EV/siRNA. Scale bars = 100 nm. (B) Size detection by NTA of EV and EV/siRNA. (C) Zeta potential detection through NTA of EV and EV/siRNA. Figure S7: siRNA loading of EVs and DP‐EVs. (A) siRNA EE detection of EVs/siRNA and DP‐EVs/siRNA complex. (B) Cell uptake efficiency of free cy3‐siRNA, EVs/cy3‐siRNA, DP‐EVs/cy3‐siRNA, and LNP/cy3‐siRNA after a 24 h incubation. Scale bars = 10 µm. Figure S8: Endosome escape of free siRNA and EVs/siRNAs. (A) Representative colocalization images at 6 and 24 h after incubation in BMDCs of early endosomes, late endosomes, and lysosomes with free cy3‐siRNA. (B) Representative colocalization images at 6 and 24 h after incubation in BMDCs of early endosomes, late endosomes, and lysosomes with EV/cy3‐siRNA. Scale bars = 5 µm. Figure S9: siRNA leakage after EVs/siRNA and DP‐EVs/siRNA complex formulation. (A) siRNA leakage detection of EVs/cy5‐siRNA and DP‐EVs/cy5‐siRNA after a 24‐h incubation with mouse serum. (B) Cell uptake efficiency of EVs/cy5‐siRNA and DP‐EVs/cy5‐siRNA after a 24‐h incubation with mouse serum in BMDCs and E.G7‐OVA. (C) PD‐L1 knockdown ability detection of EVs/cy5‐siRNA and DP‐EVs/cy5‐siRNA after a 24‐h incubation with mouse serum in BMDCs. (D) PD‐L1 knockdown ability detection of EVs/cy5‐siRNA and DP‐EVs/cy5‐siRNA after a 24‐h incubation with mouse serum in E.G7‐OVA. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Figure S10: Antitumor efficacy of DP‐EVs/siPD‐L1 in E.G7‐OVA subcutaneous tumor mouse model. (A) The timeline process of the experimental scheme. (B) Individual tumor growth curves for E.G7‐OVA tumors in mice after different treatments. (C) Average tumor growth curves for E.G7‐OVA tumors in mice after different treatments. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. Survival curves were obtained using the Kaplan–Meier method and compared by the log‐rank test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Figure S11: In vivo safety evaluation of different treatments. (A) Body weight of mice with E.G7‐OVA tumors after various treatments. (B) H&E analysis of major organs of mice after various treatments. Scale bars = 100 µm. Figure S12: The gating principle of flow cytometry for the detection of TME reprogramming in E.G7‐OVA tumors and immune response activation in lymph nodes and spleen. (A) Detection of DCs (FVS780^−^ CD45^+^ CD11c^+^), mature DCs (FVS780^−^ CD45^+^ CD11c^+^ CD86^+^MHCII^+^) and the expression of DCs (FVS780^−^ CD45^+^ CD11c^+^ PD‐L1) in lymph nodes and spleens. (B) Detection of CTLs (FVS780^−^ CD3^+^ CD8^+^ IFN‐γ^+^), Treg (FVS780^−^ CD3^+^ CD4^+^ CD25^+^ Foxp3^+^) mature DCs (FVS780^−^ CD45^+^ CD11c^+^ CD86^+^MHCII^+^), and the expression of PD‐L1 in nonimmune cells (FVS780^−^ CD45^−^ PD‐L1) in E.G7‐OVA tumors, lymph nodes, and spleens. (C) Detection of DCs (FVS780^−^ CD45^+^ CD11c^+^), mature DCs (FVS780^−^ CD45^+^ CD11c^+^ CD86^+^ MHCII^+^), M1 macrophage (FVS780^−^ CD45^+^ CD11b^+^ F4/80^+^ CD86^+^), and M2 macrophage (FVS780^−^ CD45^+^ CD11b^+^ F4/80^+^ CD206^+^) in E.G7‐OVA tumors. Figure S13: Flow cytometry detection of T cells in spleens after different treatments. (A) t‐SNE results of T cells in spleen after different treatments. (B) Flow cytometry detection of CD8^+^ IFN‐γ^+^ cells in spleen after different treatments. (C) Flow cytometry detection of Treg cells in spleen after different treatments. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Figure S14: The tumor suppression and macrophage polarization effects of DP‐EV are associated with miR‐451a and miR‐142a‐5p. (A) Heatmap of the miRNAs with different expression in DP‐EVs comparing to EVs. n = 3. (B) Detection of related miRNAs expression in DP‐EVs and EVs by qPCR. (C) Detection of EMT‐related markers (E‐cadherin, N‐cadherin, and Vimentin) at the protein levels of in E.G7‐OVA after different treatment. (D) Flow cytometry and qPCR detection of the CD206 (a marker of M2‐like macrophage) expression in macrophage after different treatments. (E) More miRNAs with different expressions and their functions. All values presented in this figure are expressed as the mean ± s.d. Statistical analysis was performed using one‐way ANOVA with Dunnett's multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
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