Nanomotor‐Driven Extracellular Vesicles With Effective Tissue Penetration for Targeted Therapy of Primary Ovarian Insufficiency
Yaoqin Mu, Jiachen Wu, Mingwei Lv, Xinyi Zhang, Yinhua Song, Jihui Ai, Ding Ma, Kezhen Li
TL;DR
A new injectable hydrogel with nanomotor-driven extracellular vesicles improves ovarian function in mice with chemotherapy-induced infertility.
Contribution
An injectable hydrogel with nanomotor-driven extracellular vesicles for targeted ovarian therapy is introduced.
Findings
The nanomotor-driven extracellular vesicles penetrate ovarian tissue and restore ovarian function in POI mice.
The therapy improves endocrine and reproductive functions by balancing oxidative stress and promoting angiogenesis.
The hydrogel enhances EV stability and retention at the ovarian site.
Abstract
Stem cells and their derived extracellular vesicles (EVs) offer hope for functional reconstruction and fertility in premature ovarian insufficiency (POI) caused by gonadotoxic drugs. However, the clinical application of EVs is impeded by their instability, limited tissue penetration, and lack of targeted delivery to ovarian injury sites. Here, we introduce an injectable hydrogel loaded with nanomotor‐driven EVs (LEVs‐Gel) for targeted POI therapy. In the POI ovarian microenvironment, reactive oxygen species (ROS) and inducible nitric oxide synthase (iNOS) are highly expressed, acting as chemoattractants to promote the chemotactic behavior of nanomotors LEVs. The LEVs are loaded in a thermosensitive hydrogel to enhance the retention and stability of EVs at the ovary. Following local injection of LEVs‐Gel into the ovaries of POI mice, the nanomotors LEVs effectively penetrated the…
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FIGURE 8- —National Key Technology Research and Development Program of China
- —National Clinical Research Center of Obstetrics and Gynecology
- —Postdoctoral Fellowship Program of CPSF
- —Major Program of the Natural Science Foundation of Hubei Province
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TopicsExtracellular vesicles in disease · Reproductive Biology and Fertility · Micro and Nano Robotics
Introduction
1
Gonadotoxic drugs, particularly chemotherapy agents like cisplatin, can cause significant damage to the ovaries, posing a major concern for the quality of life of patients post‐treatment. This issue is especially pronounced in the context of premature ovarian insufficiency (POI), which can lead to complications such as infertility, perimenopausal symptoms, and osteoporosis. Evidence indicates that women who have undergone cancer treatment exhibit a 38% reduction in pregnancy likelihood compared to the general population (Guo et al. 2024; van Dorp et al. 2018). Chemotherapy‐induced reproductive and endocrine dysfunctions arise from direct DNA damage, excessive reactive oxygen species (ROS) generation, and mitochondrial dysfunction (Dinc et al. 2023; Mentese et al., 2022; Hsueh et al. 2015; Wang et al. 2022). Agents such as paclitaxel and cisplatin substantially increase ROS levels, compromising cellular antioxidant capacity and leading to follicular atresia, premature activation of primordial follicles, and rapid depletion of the follicle pool (Zhang et al. 2023). Furthermore, chemotherapy‐associated endothelial damage and impaired angiogenesis disrupt ovarian blood supply, exacerbating follicular atresia and ovulatory dysfunction, ultimately culminating in ovarian failure and infertility (Devos et al. 2023, Zhou et al. 2021, Abdelzaher et al. 2021). Current therapeutic approaches for POI, including free radical scavengers (Ayazoglu Demir et al. 2023), angiogenic factors (Chen et al. 2023), senolytic therapies (Secomandi et al. 2022), and immunomodulators (Li et al. 2023), have demonstrated some efficacy in enhancing ovarian function. However, given the complex pathophysiology of POI, there is an urgent need for strategies that can address multiple aspects of the disease.
Considering the safety concerns surrounding mesenchymal stem cells (MSCs) therapy, MSCs‐derived extracellular vesicles (EVs) have emerged as a promising cell‐free therapeutic alternative for POI. Previous studies have shown that MSCs‐EVs can enhance granulosa cells (GCs) survival, restore follicle count, and mitigate chemotherapy‐induced reproductive toxicity (Huang et al. 2018, Yang et al. 2019, Zhang et al. 2021). However, the high ROS levels in the ovarian microenvironment post‐chemotherapy can compromise the bioactivity of EVs, as oxidative stress damages the proteins and lipids within EVs, altering their functional properties (Zhang et al. 2023; Chiaradia et al. 2021; Yarana et al. 2017; Benedikter et al. 2018). Additionally, the short half‐life of EVs (4–24 h) limits their therapeutic efficacy, particularly for the sustained administration required for POI recovery. Various carriers such as hydrogel and regenerative patches, have been explored to enhance EVs retention (Fan et al. 2024; Zhang et al. 2023). These carriers typically release EVs slowly and require penetration through the ovarian epithelium and tunica albuginea (a connective tissue layer) to reach the follicle‐rich cortex or deeper medulla, while limiting their concentration and effectiveness at the disease site (Thanuja et al. 2018; Wang et al. 2022). Obviously, the low concentration, passive diffusivity and poor tissue penetration of EVs significantly reduce their therapeutic efficiency. Therefore, it is essential to endow EVs with autonomous movement capabilities to solve the above problems. Attempts to guide nanoparticles through physiological barriers using external forces like magnetic, electrical, or optical energy have been explored; however, the ovary, being a deep‐seated abdominal organ, renders these external driving forces ineffective. Micro/nano motors with self‐propulsion capabilities driven by chemical or physical energy sources have shown great potential in enhancing barrier penetration and targeted drug delivery (Huang et al. 2025). These active movement systems promotes stronger tissue interactions and improved delivery efficiency across physiological barriers such as the brain, thymus, mucosa, testes, and placenta (Huang et al. 2025). Specifically, chemically driven nanomotors generate propulsion by reacting with the surrounding environment (e.g. hydrogen peroxide, glucose, urea, or other physiological fluids) (Wan et al. 2019), providing new possibilities for EV application in POI treatment. Thus, designing nanomotor‐based EVs capable of actively targeting and penetrating the ovarian cortex and medulla in POI is both challenging and of significant importance.
To address this challenge, we propose an injectable hydrogel loaded with nanomotor‐driven MSCs‐EVs for treating POI in mouse. As illustrated in the schematic (Figure 1A), we first construct nitric oxide(NO)‐driven nanomotor EVs (LEVs) by modifying the surface of EVs membranes derived from human umbilical cord MSCs with L‐Arginine (L‐Arg) derivative monomers. Concurrently, we synthesize a temperature‐responsive hydrogel composed of chitosan (CS)‐gelatin‐β‐glycerol (β‐GP) to serve as the LEVs carrier. By loading LEVs within the hydrogel via electrostatic interactions, we extend their local retention time in the ovaries. After ovarian intrabursae injection of LEVs‐Gel in chemotherapy‐induced POI mouse, the hydrogel gradually degrades, releasing LEVs (Figure 1B). Due to the high specific affinity of L‐Arg for NOS/ROS (Wan et al. 2019; Tan et al. 2025; Li et al. 2023; Wu et al. 2024; Chen et al. 2023), L‐Arg was catalyzed into citrulline and NO by elevated NOS/ROS levels in POI ovaries. Then NOS/ROS as chemotactic agents guide LEVs towards the ovarian cortex and medulla. The generated NO acts as a propellant, promoting active penetration into the targeted ovarian tissue. Additionally, this nanomotor does not produce any toxic byproducts, and both reactants and products are beneficial to the body (Wan et al. 2019). LEVs also balance oxidative‐antioxidative stress by scavenging ROS and NOS in POI ovaries, and the generated NO aids in ovarian angiogenesis (Wang et al. 2024). Ultimately, the multifunctional LEVs‐Gel significantly improved the ovarian function and reproductive capacity of POI mouse.
Schematic design and therapeutic mechanism of LEVs‐gel in POI treatment. (A) EVs were extracted from human umbilical cord mesenchymal stem cells (HuMSCs) and subsequently modified with cholesterol‐l‐arginine to create nanomotors (LEVs). These LEVs were incorporated into a temperature‐sensitive hydrogel composed of CS, gelatin, and β‐GP through cross‐linking, forming an injectable LEVs‐Gel. (B) The LEVs‐Gel was administered into the ovarian bursae of mice with POI. Post‐injection, the LEVs were gradually released from the hydrogel. In the POI ovarian microenvironment, LEVs are catalyzed by NOS/ROS to produce citrulline and NO. Elevated levels of NOS/ROS acted as chemoattractants, directing the nanomotors LEVs penetrate the epithelial layer and the tunica albuginea towards the ovarian cortex and medulla. The generated NO served as a driving force, enhancing the penetration into ovarian tissues and targeting sites of oxidative damage. Ultimately, the LEVs‐Gel treatment restored ovarian function in POI mice by maintaining oxidative‐antioxidative balance, inhibiting apoptosis, and promoting angiogenesis.
Results and Discussion
2
Synthesis and Characterization of LEVs Nanomotors
2.1
To synthesize NO‐driven LEVs, we strategically modified the membrane surface of EVs with L‐Arg. This process involved two primary steps: the synthesis and purification of cholesterol‐l‐Arg (CLR) (Figure S1A), followed by the incorporation of CLR into small EVs (diameter <200 nm) through physical stirring at 37°C to ensure uniform embedding into the lipid bilayer membrane. Subsequent purification yielded the final LEVs (Figure 2A). This gentle modification technique preserves the structural integrity and bioactivity of the EV membrane, which is essential for maintaining its stability and functionality. Comprehensive characterization of the purified LEVs was performed. Transmission electron microscopy (TEM) images revealed the typical cup‐shaped morphology of both EVs and LEVs (Figure 2B). Western blotting (Wb) analysis confirmed that both EVs and LEVs expressed exosomal markers, including CD9, CD63, CD81, and TSG101, while showing an absence of Calnexin expression, using human umbilical mesenchymal stem cells (HUMSCs) as a reference (Figure 2C). Nanoparticle tracking analysis (NTA) demonstrated an increase in particle size from approximately 148 nm to 168 nm and a decrease in zeta potential from ‐9.11 to ‐17.77 mV following the CLR modification of the EV membrane (Figure 2D, E). The increased negative charge of LEVs was attributed to the exposure of the carboxyl groups of L‐Arg on the surface of the compound. Fourier‐transform infrared spectroscopy (FTIR) spectra of LEVs showed the characteristic peaks of L‐Arg, a carbonyl peak at 1600–1700 cm^−1^ assigned as C = N bond) (Figure 2F). To further confirm the successful incorporation of CLR into the EV membranes, CLR was labeled with FITC (green) and EVs were labeled with Dil (red). Fluorescence colocalization studies demonstrated that LEVs exhibited both red and green fluorescence, verifying the successful integration of CLR into the EV membranes (Figure 2G). These comprehensive findings conclusively validate the successful synthesis and precise characterization of LEVs.
*Characterizations of LEVs nanomotors’ structure and motion behavior. (A) Schematic of the preparation and working principle of LEVs nanomotors. (B) TEM images of EVs and LEVs. Scale bar, 200 nm. (C) Expression analysis of biomarkers CD9, CD63, CD81, and TSG101 in HUMSCs, EVs, and LEVs. (D) Size distribution of EVs and LEVs. (E) Zeta potential of EVs and LEVs. (F) Fourier transform infrared spectroscopy (FTIR) of L‐Arg, EVs, and LEVs. (G) Confocal fluorescence images of EVs and LEVs, with FITC in green and Dil in red, scale bar: 100 µm. (H) Representative images showing intracellular ROS levels in EVs and LEVs, scale bar: 100 µm. (I) Mean fluorescence intensity (MFI) analysis of DCFH‐DA. (J) Relative activity analysis of NOS. (K) Analysis of relative nitric oxide (NO) production by EVs and LEVs at 6, 12, 24, and 48 h in the POI cell model. (L and M) Normalized movement trajectories of EVs (L) and LEVs (M) in POI cell model (Video S1 and S2, Supporting Information, n = 10). (N) Velocity analysis of EVs and LEVs. (O) Schematic illustration of the Y‐shaped channel. (I) Dil‐labeled LEVs in reservoir; (II) agarose gel containing KGN cell lysate; (III)agarose gel containing POI‐KGN cell lysate. (P) Representative images of LEVs in reservoirs (II) and (III) of the Y‐shaped channel. (Q) Fluorescence intensity (FI) analysis of LEVs in reservoirs. (R) Confocal fluorescence images of cellular uptake of EVs or LEVs after co‐incubation for 60 and 120 min. Scale bar:10 µm (*p < 0.05, **p < 0.01,***p < 0.001 and ***p < 0.0001).
Assessment of Antioxidative, Autonomous Motion, and Chemotactic Properties of LEVs Nanomotors
2.2
Targeted therapeutic delivery aims to precisely direct drugs to pathological sites, significantly enhancing efficacy while minimizing adverse effects. Traditional EVs rely on passive diffusion driven by concentration gradients, often resulting in limited ability to penetrate physiological barriers. In contrast, micro/nanoscale motors with self‐propulsion capabilities exhibit enhanced barrier penetration, thus improving targeting efficiency. Inspired by the endogenous conversion of L‐Arg to NO by NOS or ROS, which chemotactically guides NO to sites of inflammation or injury (Wan et al. 2019, Li et al. 2023), we designed NO‐driven LEVs to enhance EVs penetration into the deeper cortical and medullary regions of POI ovaries.
To validate the properties of LEVs nanomotors, we systematically evaluated their ability to eliminate ROS/NOS, NO production, motility, and chemotaxis. Oxidative stress is a hallmark of chemotherapy‐induced POI, characterized by significantly elevated ROS and NOS levels in cisplatin‐induced POI cell models (Figure 2H). Administration of 100 µg/mL LEVs significantly reduced ROS and NOS levels (Figure 2H‐J). Moreover, LEVs exhibited superior antioxidative properties compared to conventional EVs, suggesting that L‐Arg modification on the LEVs membrane facilitates rapid ROS and NOS scavenging. This high affinity with ROS/NOS provides the foundation for the production of NO (Wan et al. 2019). In the POI cellular environment, LEVs consistently released NO for up to 48 h (Figure 2K), providing the fundamental propulsion mechanism for their nanomotor characteristics. Tracking the motion of EVs and LEVs in POI cell culture medium revealed that while EVs exhibited typical Brownian motion (Figure 2L, Video 1), LEVs driven by nanomotors move in different directions and displayed parabolic trajectories indicative of autonomous movement (Figure 2M, Video 2). The movement speed of LEVs was significantly higher than that of EVs, likely driven by NO (Figure 2N). These findings suggest that L‐Arg on the LEVs membrane is catalyzed by the high NOS/ROS levels in the POI microenvironment to produce NO, fueling LEVs motion. To assess the ROS/NOS chemotactic behavior of LEVs, we designed a Y‐shaped channel (Figure 2O). Dil‐labeled LEVs were loaded into reservoir (I), with gels containing KGN and POI‐KGN cell lysates placed in reservoirs (II) and (III), respectively, to create chemotactic gradients of ROS and NOS (Figure S1B). Over time, fluorescence intensity in reservoir (III) significantly increased at the 60‐min and the 90‐min mark, while reservoir (II) remained unchanged (Figure 2P, 2Q). Control experiments with EVs instead of LEVs in reservoir (I) showed no significant fluorescence change within 90 min (Figure S1C). These results confirm that LEVs actively migrate towards target sites driven by ROS/NOS gradients. Furthermore, effective targeting necessitates cellular uptake of LEVs. Confocal microscopy revealed that Dil‐labeled EVs/LEVs were taken up by KGN/POI‐KGN cells within 1–2 h, with LEVs entering POI‐KGN cells more rapidly, indicating that autonomous motion enhances LEVs uptake (Figure 2R).
In conclusion, in vitro assessment of LEVs antioxidative capacity, autonomous motility, and chemotactic behavior establishes that LEVs possess nanomotor properties, enabling their autonomous and chemotactic movement within the NOS/ROS gradient of the POI microenvironment.
Synthesis and Characterization of LEVs‐Gel
2.3
Next, we synthesized a CS/β‐GP/gelatin hydrogel as a delivery vehicle for LEV nanomotors, integrating these nanomotor‐type LEVs into the hydrogel to enhance therapeutic efficacy for clinical POI.
The thermoresponsive behavior of the CS/β‐GP/gelatin hydrogel is illustrated in Figure 3A. At 25°C, the CS/β‐GP/gelatin exists as a flowable hydrogel precursor. Upon increasing the temperature, hydrophobic interactions among CS, β‐GP, and gelatin intensify, leading to gelation. Previous studies have demonstrated that the inclusion of gelatin effectively shortens the gelation time by accelerating crosslinking between CS and β‐GP (Cheng et al. 2011, Liu et al. 2025). To synthesize a rapidly gelating CS/β‐GP/gelatin hydrogel, we optimized the gelatin concentration (0.3%, 0.6%, and 0.9%). Scanning electron microscopy (SEM) revealed that all hydrogel formulations developed a porous three‐dimensional network structure with uniform pore distribution, and the average pore size decreased with increasing gelatin concentration (ranging from 7 to 0.5 µm) (Figures 3B, S2A and S2B). We then loaded 500 µg of LEVs into each hydrogel at concentrations of 0.3%, 0.6%, and 0.9% gelatin, with hydrogels without LEVs serving as controls. The net amount of LEVs released was quantified by BCA assay on days 1, 2, 3, 5, and 7. Then, we found that the 0.3% and 0.6% gelatin groups exhibited daily net releases of approximately 150–200 µg and 100–150 µg on days 1 and 2, respectively, with a gradual decrease from day 3. In contrast, the 0.9% gelatin group maintained a near‐constant daily net release of around 50 µg, yielding a flatter overall release profile. These results are consistent with SEM analyses showing that higher gelatin concentrations reduce hydrogel pore size and enhance LEVs retention. At 25°C, all formulations exhibited good fluidity; however, at 37°C, hydrogel containing 0.3%, 0.6%, and 0.9% gelatin transitioned to gel states within 180 s, 180 s, and 150 s, respectively (Figure 3C, S2C and S2D). Considering both LEVs retention and gelation time, we selected the hydrogel with 0.9% gelatin for further experiments. Additionally, the hydrogel remains injectable at room temperature (Figure 3D). Rheological assessments revealed that the storage modulus (G') of the hydrogels exceeded the loss modulus (G'') at 4°C, 25°C, and notably at 37°C, confirming the thermoresponsive gelation properties of the CS/β‐GP/gelatin hydrogel (Figure 3E). Tensile tests indicated that the hydrogel could withstand elongation up to 4.8 times its original length and a maximum tensile force of approximately 2.49 MPa, demonstrating excellent mechanical properties and stability (Figure 3F).
Synthesis and characterization of LEVs‐Gel. (A) Schematic illustration of temperature‐responsive hydrogel. (B) Scanning electron microscopy (SEM) image of hydrogel containing 0.9% gelatin and net release of LEVs over 7 days. The initial content of LEVs is 500 µg. Scale bar:10 µm. (C) Image of the gel formation of CS/β‐GP/gelatin solution with 0.9% gelatin. (D) Injectable property of CS/β‐GP/gelatin hydrogel. (E) Storage modulus (G′) and loss modulus (G′′) of hydrogel at 4°C, 25°C and 37°C. (F) Tensile tests of hydrogel. (G) Illustration of LEVs loading into the hydrogel via electrostatic interactions. (H) Zeta potential analysis of EVs, LEVs and LEVs‐Gel. (I) Confocal fluorescence images of LEVs loaded in the hydrogel. (J) The accumulative release of LEVs from the hydrogel containing 0.9% gelatin over 21 days. The initial content of LEVs is 1000 µg. (K) Assessment of cell survival with hydrogel loaded with varying amounts of LEVs.
Given the presence of abundant amino and carboxyl groups in the CS/β‐GP/gelatin hydrogel and the negatively charged LEVs, we hypothesized that LEVs could bind to the hydrogel through electrostatic interactions (Figure 3G). To verify this, we lyophilized the hydrogel into particles and measured the zeta potential. The hydrogel exhibited a mean charge of +12.60 mV (Figure 3H), suggesting that LEVs, with their negative charge, could be effectively adsorbed by the positively charged hydrogel, facilitating stable LEVs loading and prolonged retention time. Confocal microscopy confirmed that Dil‐labeled LEVs were uniformly distributed within the hydrogel (Figure 3I). To investigate the in vitro release kinetics of LEVs from LEVs‐Gel, we incubated LEVs‐Gel containing 1000 µg of LEVs in Transwell chambers at 37°C in PBS. The release profile showed an initial burst followed by a sustained release, with over 70% cumulative release over 24 days (Figures 3J and S2E). These results indicate that the hydrogel not only evenly and stably incorporates LEVs but also ensures their sustained release. To assess the biocompatibility of the LEVs‐Gel system, we conducted a series of in vitro assays. Different amounts of LEVs were loaded into the hydrogel to form several experimental groups. The CCK‐8 assay results demonstrated that LEVs‐Gel did not adversely impact cell proliferation (Figure 3K). Based on these findings, we selected LEVs‐Gel loaded 200 µg LEVs for subsequent experiments. Further evaluation using live/dead staining (calcein‐AM/PI) and Ki67 immunofluorescence revealed that LEVs‐Gel with 200 µg LEVs did not compromise cell viability or proliferation (Figure S3A–D). Flow cytometric analysis of apoptosis further confirmed that, in a POI cell model, LEVs‐Gel significantly reduced cell apoptosis rates (Figure S3E). These comprehensive assessments indicate that LEVs‐Gel exhibits excellent biocompatibility. Collectively, our findings affirm the successful preparation and characterization of the LEVs‐Gel system, highlighting its promising potential as a therapeutic platform for the treatment of POI.
LEVs‐Gel Enhanced Retention and Tissue Penetration of EVs in POI Mouse Ovaries
2.4
To assess the retention and tissue penetration of LEVs‐Gel in mouse ovaries, we developed a POI model through intraperitoneal injection of cisplatin. Based on the above results of the biocompatibility and in vitro release experiments of LEVs, we loaded LEVs in the hydrogel and administered a single ovarian injection. The hydrogel provided prolonged and slow release at low concentrations in vitro, thereby reducing the risk of cytotoxicity associated with the exposure to high‐concentration EVs. Furthermore, previous studies have shown that 200 µg EVs significantly increased the number of ovarian follicles, promoted the proliferation of granulosa cells, and inhibited the apoptosis of granulosa cells damaged by chemotherapy, as well as those in the cultured ovaries and the ovaries of mice (Cao et al. 2023). Therefore, we selected a 200 µg EVs/LEVs dose for in vivo studies in POI mice. We administered Dil‐labeled 200 µg EVs, 200 µg LEVs, and LEVs‐Gel loaded with 200 µg LEVs into the ovarian bursa of POI mouse. Utilizing a real‐time imaging system, we tracked the fluorescence signals of EVs at multiple time points: 0, 12, 24, 48, 72 h, 7, 14, and 21 days post‐injection in excised ovaries (Figure 4A, D). The EVs group exhibited rapid leakage towards the fallopian tubes and uterus, retaining approximately 13% of the EVs at 48 h post‐injection, which further degraded to 5% by 72 h. This pattern is consistent with previous reports that local ovarian injection of EVs lead to swift clearance and degradation. Compared with the EVs group, LEVs group demonstrated a retention rate of about 19% at 48 h, dropping to 13% at 72 h, and was undetectable by day 7, highlighting the rapid degradation of LEVs in vivo. Obviously, both the EVs group and the LEVs group have the problem of short retention time in vivo, which fails to meet the long‐term therapeutic requirements for ovarian treatment. Compared with the LEVs group, due to the electrostatic interactions between the hydrogel and LEVs combined with the hydrogel's slow‐release capabilities, LEVs‐Gel group exhibited a biphasic release profile characterized by an initial rapid release followed by sustained release. At 48 and 72 h post‐injection, the LEVs‐Gel group retained 3.3‐fold and 4.3‐fold more compared to the LEVs group. By day 7, approximately 49% of the EVs remained in LEVs‐Gel group, with about 16% still present at day 21 (Figure 4A, D). These findings demonstrate that LEVs‐Gel significantly prolongs the local retention time of LEVs in the ovaries, thereby enhancing their therapeutic potential.
*Retention and tissue penetration of EVs in LEVs‐Gel within ovaries. (A) Fluorescence images of mouse uterus and ovaries in various groups at different time points. (B) Fluorescence images of whole ovarian tissue and longitudinal sections. (C) Fluorescence images of whole ovarian tissue and horizontal slice sections. (D) Analysis of EVs retention in various groups over time. (E) Fluorescence intensity analysis of longitudinal slice section. (F) Fluorescence intensity analysis of horizontal slice section. (ns: p > 0.05,*p < 0.05, **p < 0.01,***p < 0.001 and ***p < 0.0001).
The ovarian tissue is composed of an outer epithelial layer, the tunica albuginea (a connective tissue layer), a cortex abundant in follicles, and a medulla rich in vasculature. Effective targeted therapy of EVs in the ovary requires traversing these physiological barriers to reach the cortex and medulla, where therapeutic action is most needed. Our in vitro experiments have confirmed the active motility of nanomotor LEVs. And then we further investigated the permeability of nanomotor LEVs within mouse ovarian tissue. Ovaries treated for 48 h were sectioned both longitudinally and transversely to evaluate the distribution and retention of EVs among varies groups. To confirm the chemotactic movement of nanomotors in LEVs‐Gel, we included a healthy Control+LEVs‐Gel where LEVs‐Gel was administered to the ovaries of healthy mice. Longitudinal sections (Figure 4B, E) revealed that in the EVs group, EVs primarily accumulated at the ovarian tissue margins, with minimal penetration into the cortex and medulla. This indicates poor tissue penetration, likely due to the challenges of crossing the ovarian epithelium and tunica albuginea. Conversely, in the LEVs group, a notable accumulation of EVs was observed in the ovarian cortex and medulla, suggesting that despite their limited retention, LEVs could effectively penetrate the ovarian tissue. Remarkably, the LEVs‐Gel group exhibited a uniform distribution of a substantial number of EVs in both the cortex and medulla, with significantly higher average fluorescence intensity compared to the other groups (all p < 0.05). This indicates that LEVs‐Gel not only enhanced the retention time of EVs but also improved their tissue permeability. By contrast, the overall red fluorescence intensity in the ovaries of the Control+LEVs‐Gel group remained high, with EVs predominantly located at the ovarian margins and sparsely present in the cortex and medulla. The high fluorescence intensity of the entire ovary might be due to EVs residing on the ovarian surface or within the surrounding adipose tissue. Therefore, while LEVs‐Gel can extend EVs retention time, the low ROS and NOS levels in the ovaries may limit the chemotactic movement of LEVs. These observations suggest that elevated ROS and iNOS levels in POI ovaries stimulate robust NO generation by LEVs, thereby driving active motion toward deeper medullary regions. Conversely, the lower ROS/iNOS levels in healthy ovaries do not elicit sufficient NO to power nanomotor motion, resulting in predominantly passive diffusion restricted to the cortex. Transverse sections further confirmed that the tissue accumulation of EVs in the LEVs‐Gel group was significantly higher than in the other groups (all p < 0.05) (Figure 4C, F). These findings suggest that LEVs‐Gel substantially enhances both the retention and tissue penetration of EVs within the ovaries of POI mice, thereby improving the effective utilization of EVs and contributing to the long‐term recovery of ovarian function in POI.
LEVs‐Gel Promoted Recovery of Ovarian Function in POI Mouse
2.5
To further assess the therapeutic efficacy of LEVs‐Gel in a mouse model of POI, treatments were administered 21 days post‐chemotherapy, with subsequent assessments of ovarian function conducted 21 days following treatment (Figure 5A). The ovary is a critical endocrine organ in the female reproductive system, and chemotherapy can disrupt ovarian hormone levels (Rives et al. 2022). Our study revealed that LEVs‐Gel treatment showed a tendency of improvement of ovarian index compared to the POI group, although the difference was not statistically significant (p > 0.05) (Figure 5B). Estrous cycle monitoring demonstrated that both LEVs and LEVs‐Gel treatments effectively restored the physiological rhythm in POI mouse (Figures 5C and S4A). Consistent with previous studies, POI mouse exhibited decreased serum estradiol (E2) levels and elevated follicle‐stimulating hormone (FSH) and luteinizing hormone (LH) levels (Zhang et al. 2023; Shin et al. 2021). Notably, LEVs‐Gel treatment significantly increased the levels of E2 and FSH, and this therapeutic effect is more significant than that of the other groups (all p < 0.05) (Figure 5D–F). It is worth noting that after 21 days of LEVs‐Gel treatment in POI mice, the ovarian index and LH levels showed partial recovery (p > 0.05). Previous work has shown that EVs or exosomes often require a longer time frame to restore the ovarian index and LH in POI mice (Cui et al. 2025; Park et al. 2024; Zhou et al. 2024). For example, in one study using mesenchymal stem cell derived exosomes, the ovarian index and LH level recovered after 28 days of treatment and followed by a 21‐day observation (Cui et al. 2025). In our study, based on the in vivo release profile of EVs from the LEVs‐Gel, we assessed ovarian function at day 21. In future studies, we will investigate longer treatment and observation durations to determine whether further improvements in the ovarian index and LH level can be achieved.
*The effect of LEVs‐Gel treatment on ovarian function in POI. (A) Schematic illustration of LEVs‐Gel treatment plan for POI. (B) Analysis of ovarian indices from different groups. (C) Analysis of mouse estrous cycle from different groups. (D–G) Analysis of serum hormone levels from different groups: E2 (D); FSH (E); LH (F); AMH (G). (H) Representative histological images of ovarian sections from each group. (I) Quantification of ovarian follicles at different developmental stages across groups. (nc: p > 0.05, *p < 0.05, **p < 0.01,***p < 0.001 and ***p < 0.0001).
A complete reproductive function depends on maintaining an adequate ovarian reserve, where anti‐Müllerian hormone (AMH) levels and the count of ovarian follicles serve as primary indicators. As expected, chemotherapy induced a marked decline in AMH levels in POI mice. LEVs‐Gel treatment significantly elevated AMH levels compared to EVs and LEVs treatments (p < 0.05) (Figure 5G). Previous studies have demonstrated that chemotherapeutic agents cause follicular atresia and premature depletion of the follicular pool (Hsueh et al. 2015; Wang et al. 2022; Shin et al. 2021). Our results corroborate these findings, showing a significant reduction in primordial, primary, secondary, and antral follicles, alongside an increase in atretic follicles in POI mice subjected to chemotherapy (control vs. POI, all p < 0.05) (Figure 5H, I). LEVs‐Gel treatment substantially increased the number of follicles at various developmental stages and decreased the number of atretic follicles compared to the EVs and LEVs groups. Specifically, LEVs‐Gel significantly promoted the growth, development, and maturation of primordial follicles while inhibiting follicular atresia, thereby restoring ovarian reserve function (POI vs. LEVs‐Gel, all p < 0.05) (Figure 5H, I). Collectively, these results affirm that LEVs‐Gel markedly enhances the recovery of ovarian function in the POI mouse model, highlighting its potential as a promising therapeutic strategy for restoring fertility and endocrine function in chemotherapy‐induced POI.
LEVs‐Gel Promoted the Restoration of Reproductive Capacity in POI Mouse
2.6
Chemotherapy‐induced POI is not only characterized by reduced ovarian function but also poses substantial challenges to fertility (Spears et al. 2019). To address these issues, we we examined the impact of LEVs‐Gel on the reproductive capacity of POI mouse, with a focus on oocyte quality, in vitro developmental potential, and fertility outcomes. In vitro maturation (IVM) results revealed that the maturation rates in the EVs, LEVs, and LEVs‐Gel treatment groups were significantly higher compared to the POI group. Notably, the LEVs‐Gel group exhibited a pronounced increase in oocyte maturation rates (POI vs. LEVs‐Gel, P<0.05) (Figure 6A, B). Furthermore, in vitro fertilization (IVF) outcomes demonstrated that LEVs‐Gel treatment substantially enhanced both fertilization rates and blastocyst formation rates relative to the POI, EVs, and LEVs groups (POI vs. LEVs‐Gel, p < 0.05) (Figure 6C, D). The application of LEVs‐Gel in POI mice significantly increased the recruitment of mature oocytes and improved both in vitro fertilization and blastocyst formation rates. This suggests that LEVs‐Gel could potentially enhance the success rates of assisted reproductive technologies in women who have undergone chemotherapy.
*The effect of LEVs‐Gel treatment on the reproductive fertility in POI. (A) Representative images of MII oocytes in vitro maturated from different groups. (B) Analysis of the MII oocyte maturation rate. (C) Representative images of 2‐cell stage embryos and blastocysts in vitro fertilized oocytes from different groups. (D) Analysis of 2‐cell stage embryos and blastocysts formation rates. (E) Schematic illustration of reproductive fertility testing post LEVs‐Gel treatment. (F) Representative images of first generation baby mouse. (G–O) Reproductive capacity of the parental mice (G–J) and first‐generation mice (L–O), including fetal mortality (G and L), average live birth count (H and M), sex ratio (I and N), and viability (J and O). (K) Body weight measurement of the first‐generation mice. (nc: p > 0.05, *p < 0.05, **p < 0.01 and **p < 0.001).
We further evaluated the impact of LEVs‐Gel on natural fertility and offspring health in POI mouse, including the reproductive capacity of both parental and F1 generation mouse, as well as the growth and health status of the offspring (Figure 6E). Consistent with previous reports (Morgan et al. 2012), chemotherapy significantly impaired the fertility of POI mouse (NC vs. POI, p < 0.05). LEVs‐Gel treatment for 21 days significantly increased the average live birth count in POI mouse compared to the EVs and LEVs groups (POI vs. LEVs‐Gel, p < 0.05) (Figure 6F, H). Additionally, there were no significant differences in fetal mortality, sex ratio, and viability among the groups (p > 0.05) (Figure 6G, I, and J). Furthermore, the reproductive capacity of the F1 generation, including fetal mortality, average live birth count, sex ratio, and viability, did not show significant differences across all groups (Figure 6L–O) (all p > 0.05). Growth and development indicators, such as early body weight, ear opening, lower incisor eruption, eye opening, cliff avoidance test, and negative geotaxis test, also showed no significant differences among the groups (all p > 0.05) (Figures 6K and S4B–G). These results indicate that LEVs‐Gel treatment does not impair the fertility or health of the offspring, affirming its efficacy in improving post‐chemotherapy fertility in POI mouse without adverse effects on subsequent generations. In vivo biosafety of LEVs‐Gel was validated through hematoxylin and eosin (HE) staining, which revealed healthy morphology of the heart, liver, spleen, lungs, and kidneys across all groups, with no observable inflammatory cell infiltration or pathological changes (Figure S5A). Additionally, blood biochemical indices of the mice remained normal 3 months post‐transplantation, indicating no significant hepatotoxicity or nephrotoxicity (Figure S5B). These results suggested that LEVs‐Gel has no significant toxic side effects.
Our in vivo results clearly demonstrate that LEVs‐Gel effectively restores ovarian function and reproductive capacity in POI mouse, without compromising the immediate or long‐term health of the offspring. This provides strong evidence for the potential clinical application of LEVs‐Gel. Moreover, we investigated the underlying molecular mechanisms of the LEVs‐Gel system in facilitating these therapeutic effects.
RNA‐seq Analysis of LEVs‐Gels Effect on POI Mouse Ovary
2.7
To elucidate the molecular mechanisms underlying the therapeutic effects of LEVs‐Gel on POI, we conducted RNA sequencing (RNA‐seq) to analyze the ovarian transcriptome of POI mice. Total RNA was extracted from the ovaries of three groups: normal control mice (Control), untreated POI mice, and POI mice treated with LEVs‐Gel. Principal component analysis (PCA) and component correlation analysis confirmed the consistency and reliability of the RNA‐seq data across all groups (Figure 7A, F). Differential gene expression analysis revealed significant transcriptional changes. Compared to the control group, 297 genes were downregulated and 23 genes were upregulated in the POI group (Figure 7B). In contrast, relative to the POI group, treatment with LEVs‐Gel resulted in the downregulation of 257 genes and upregulation of 549 genes (Figure 7G). Heatmaps of differential gene expression across the samples further substantiated the reproducibility of these findings (Figure 7C, H).
Transcriptomics analysis in the POI mouse model. (A and F) Principal component analysis. (B and G) Differential expressed genes (DEGs) analysis. (C and H) Heatmap of DEGs. (D and I) GO enrichment analysis of DEGs covering biological process (BP), molecular function (MF) and cellular components (CC). (E and J) KEGG enrichment analysis of DGE. (A–E): Control vs. POI; (F–J): POI vs. LEVs‐Gel.
Gene Ontology (GO) functional enrichment analysis (Top 10 terms) indicated that downregulated genes in the POI group were predominantly involved in the synthesis and metabolism of critical biomolecules such as cholesterol and steroids, suggesting chemotherapy‐induced impairment of hormonal endocrine function (Figure 7D). Additionally, these downregulated genes were related to oxidoreductase activity, peroxisome function, and mitochondrial crista integrity, implying a reduction in ovarian antioxidant capacity. Conversely, LEVs‐Gel treatment upregulated genes associated with ovarian hormone endocrine function and antioxidant capacity, highlighting its restorative effects (Figure 7I). KEGG pathway enrichment analysis (Top 20 pathways) further demonstrated that downregulated genes in the POI group were involved in ovarian hormone endocrine function and peroxidase signaling pathways when compared to the control group (Figure 7E). Following LEVs‐Gel treatment, upregulated genes were primarily associated with pathways related to ovarian hormone endocrine function and antioxidant signaling (Figure 7J). These findings are consistent with our above results that LEVs‐Gel restores ovarian function in POI mice. Untreated POI mice showed reduced ovarian reserve and endocrine function, evidenced by lower AMH levels, fewer follicle counts, decreased E2, and elevated FSH and LH levels, whereas LEVs‐Gel treatment promoted recovery with increased AMH levels, higher follicle counts, elevated E2, and reduced FSH and LH levels. In addition, LEVs‐Gel treatment also activated pathways linked to angiogenesis, such as the HIF‐1 and cGMP‐PKG signaling pathways.
The Molecular Mechanism of LEVs‐Gel in Restoring Ovarian Function in POI Mouse
2.8
Oxidative stress is a pivotal mechanism driving chemotherapy‐induced ovarian dysfunction. Chemotherapeutic agents, such as cisplatin and cyclophosphamide, elevate ROS levels and inhibit antioxidant enzyme activity, thereby disrupting the oxidative‐antioxidative balance within the ovary (Guo et al. 2024, Morgan et al. 2012). This imbalance induces apoptosis and compromises oocyte quality. Based on the design principles of LEVs and RNA sequencing results, we further verified the antioxidative effects of LEVs‐Gel. Ovarian tissue lysates from different mouse groups were collected, and oxidative and antioxidative parameters were assessed. The results demonstrated that chemotherapy significantly increased oxidative stress in the ovaries of POI mice, as evidenced by elevated levels of ROS, induced iNOS, and 8‐hydroxy‐2'‐deoxyguanosine (8‐OHdG) (Figure 8A–C). Concurrently, the antioxidative capacity was markedly reduced, indicated by decreased levels of total antioxidative capacity (T‐AOC), glutathione peroxidase 1 (GPX1), and glutathione synthetase (GSS) (Figure 8D, E). LEVs‐Gel treatment notably maintained the oxidative‐antioxidative balance in the ovaries of POI mice (Figure 8F). This is consistent with our in vitro results, suggesting that elevated ROS and iNOS in POI ovaries may be neutralized by NO generated from the L‐Arg‐rich surface of LEVs, thereby mitigating oxidative stress. Additionally, chemotherapeutic agents directly induce oxidative damage to oocytes. Our findings demonstrated that ROS levels in oocytes (Figures S6A, S6B) were elevated and mitochondrial membrane potential (MMP) was reduced in POI mice (Figure S6C and S6D). However, LEVs‐Gel treatment significantly ameliorated oxidative stress in oocytes.
*Molecular mechanism underlying LEVs‐Gel‐mediated ovarian function recovery. (A–C) Analysis of oxidative stress markers in ovarian tissues from POI mice post LEVs‐Gel treatment, including ROS (A), iNOS (B) and 8‐OHdG (C) using ELISA kits. (D) Analysis of total antioxidant capacity (T‐AOC) in ovarian tissues via biochemical assays. (E) Western blot analysis of antioxidant proteins GPX1 and GSS in ovarian tissues from each group. (F) Changes in the balance of oxidation and antioxidation within the ovaries of POI mice before and after LEVs‐Gel treatment. (G and H) Representative immunohistochemical staining images of H2AX (G) and cleaved caspase‐3 (H) in ovarian tissues from different groups. (I and J) Quantitative analysis of H2AX and cleaved caspase‐3 positive area fraction. (K) Wb analysis of signaling pathway proteins including MEK1/2, p‐MEK1/2, ERK1/2, p‐MEK1/2, VEGFA, HIF‐1α and tubulin. (L) Representative immunohistochemical staining images and quantitative analysis of CD34 expression in ovarian tissues. (M) Schematic depiction of the molecular mechanisms by which LEVs‐Gel promote angiogenesis in the ovaries of POI mice. (nc: p > 0.05, *p < 0.05, **p < 0.01,***p < 0.001 and ***p < 0.0001).
We further evaluated DNA damage and apoptosis in ovarian cells by examining the phosphorylation of H2AX and the expression of cleaved caspase‐3. Phosphorylated H2AX (Figure 8G, I) and cleaved caspase‐3 (Figure 8H, J) were predominantly localized in GCs. Compared to the POI group, treatment with LEVs‐Gel significantly reduced H2AX phosphorylation and the proportion of cleaved caspase‐3‐positive GCs (all P<0.05), suggesting that LEVs‐Gel effectively mitigates apoptosis in the ovaries of POI mouse.
Chemotherapy‐induced ovarian dysfunction is also attributed to disrupted angiogenesis. Chemotherapeutic agents impair ovarian vascular density, integrity, and function, which are essential for follicular development (Guo et al. 2024). RNA‐seq results indicated that LEVs‐Gel could enhance the formation of new blood vessels in the ovary. We further verified at the protein level whether LEVs‐Gel could promote angiogenesis in ovarian microvessels and activate related angiogenesis pathways. Wb analysis revealed that LEVs‐Gel treatment significantly increased the phosphorylation levels of MEK1/2 and ERK1/2 and the expression of HIF‐1α and vascular endothelial growth factor A (VEGFA), compared to the POI group (p < 0.05) (Figure 8K). These results indicated that LEVs‐Gel activated the activity of MEK1/2, ERK1/2, upregulated HIF‐1α expression, and ultimately increased VEGFA expression. In particular, the expression of VEGFA protein directly stimulates angiogenesis (Li et al. 2025; Han et al. 2025). Microvessel density was calculated by CD34 immunohistochemistry. Immunohistochemical analysis of CD34 distribution in the ovaries showed that CD34 was mainly expressed around follicles and within the ovarian stroma (Figure 8L). Chemotherapy markedly reduced microvessel density in the ovaries of POI mice, whereas LEVs‐Gel treatment significantly increased microvessel density (Figure 8L). Therefore, LEVs‐Gel may have promoted angiogenesis in POI mice by regulating the HIF‐1α/VEGFA signaling pathway (Figure 8M).
To sum up, LEVs‐Gel enhances overall ovarian function in POI mice by maintaining the oxidative‐antioxidative balance within the ovarian microenvironment, inhibiting cell apoptosis, and promoting ovarian angiogenesis.
Conclusion
3
In summary, we developed an injectable hydrogel loaded with nanomotor‐driven LEVs (LEVs‐Gel) for the treatment of POI mouse. In vitro experients demonstrated that elevated levels of ROS and NOS in the POI ovarian microenvironment as chemotactic agents, guiding LEVs’ active motility. LEVs were electrostatically loaded into a CS/β‐GP/gelatin hydrogel, which is temperature‐responsive, enhancing its injectability and prolonging the release time of EVs. In vivo results revealed that LEVs‐Gel could be retained within the ovaries for up to 21 days, and the controlled release of LEVs allowed them to penetrate the epithelial layer and the tunica albuginea of the POI ovaries, reaching the ovarian cortex and medulla to exert their effects, thereby significantly improving the utilization efficiency of LEVs. Moreover, the local injection of LEVs‐Gel into the ovarian bursa of POI mouse promoted the restoration of ovarian function and fertility. Mechanistically, we demonstrated that LEVs‐Gel maintained the oxidative‐antioxidative balance in the ovaries of POI mouse, reduced granulosa cell apoptosis, and minimized oocyte oxidative damage. Additionally, it enhanced ovarian angiogenesis by modulating the HIF‐1α/VEGFA signaling pathway, ultimately improving ovarian function.
Methods
4
Preparation, Characterization, and Fluorescent Label of LEVs
4.1
EVs was quantified using the bicinchoninic acid (BCA) assay. EVs were then mixed with CLR at a mass ratio of 1:5. The mixture was incubated in sterile centrifuge tubes at 37°C with rotation at 500 rpm for 2 h. Unbound CLR was removed using a purification column, and the LEVs were concentrated using a centrifugal concentrator. FTIR spectroscopy was employed to detect specific absorption peaks on the modified LEV membranes. Additional characterizations were performed as previously described. For fluorescent labeling, CLR was mixed with FITC dye and incubated at 37°C for 6 h to impart green fluorescence to CLR. EVs/LEVs suspensions were then stained with DiI dye at 37°C for 30 min to impart red fluorescence to the membranes.
NO Production by Nanomotors in POI‐KGNs
4.2
NO production in POI‐KGNs was assessed using a NO detection kit. POI‐KGNs (1 × 10^5^ cells/mL) were seeded in 96‐well plates and incubated for 12 h. The medium was then replaced, and the cells were treated with 100 µg/mL EVs or LEVs. Culture supernatants were collected at 6, 12, 24, and 48 h post‐treatment, and NO concentrations were measured according to the NO detection kit protocol. Collect the liquid from each site within the channel, and use the ELISA kit to detect the concentrations of ROS and NOS.
Chemotactic Behavior of Nanomotors in a Y‐Shaped Channel
4.3
According to the previous research methods, we employed a Y‐shaped PDMS channel with a main channel length of 1 cm and width of 0.4 cm, and branch channels each 0.7 cm in length and 0.3 cm in width. We collected KGN cells from control and POI groups and lysed them using ultrasonic disruption to extract their contents. These extracts were then mixed with equal volumes of agarose solution in reservoirs (ii) and (iii). The Y‐shaped channel was filled with 300 µL of PBS, and 60 µL of Dil‐labeled LEVs or EVs solution was added to reservoir (i). Fluorescent images of reservoirs (ii) and (iii) were captured at predetermined time points using an inverted fluorescence microscope.
Trajectories of LEVs
4.4
Supernatants from POI‐KGN cells were collected and centrifuged at 120,000 × g for 15 min to remove cellular debris. One microliter of Dil‐labeled EVs/LEVs was added to 10 µL of the aforementioned cell culture supernatant and mixed thoroughly. The motion of EVs/LEVs was observed and recorded under a confocal microscope for no less than 10 s. Trajectories and velocities of EVs and LEVs were analyzed using Fiji image software.
Distribution and in Vitro Release Profile of LEVs in the LEVs‐Gel System
4.5
Dil‐labeled LEVs were loaded into the temperature‐responsive Hydrogel. The LEVs‐Gel was added to the center of a confocal dish and incubated at 37°C until gelation. The distribution of LEVs was observed using a confocal microscope. For in vitro release testing, We first synthesized thermoresponsive hydrogels at concentrations of 0.3%, 0.6%, and 0.9% gelatin. We then loaded 500 µg of LEVs into each hydrogel, with hydrogels without LEVs serving as controls. The net amount of LEVs released was quantified by BCA assay on days 1, 2, 3, 5, and 7. After determining the concentration of gelatin, the release amount of 1000 µg of LEVs at different time points (Day1, 2, 3, 6, 9, 12, 15, 18, and 21) in the hydrogel was measured.
In Vivo Distribution and Retention of LEVs‐Gel
4.6
Dil‐labeled EVs/LEVs were injected into the ovarian bursa of POI mice in the following groups: 10 µL of 200 µg EVs, 10 µL of 200 µg LEVs, and 10 µL of LEVs‐Gel loaded with 200 µg LEVs. A blank control group received 10 µL of normal saline. Mice were euthanized at 0, 12, 24, 48, 72 h, 7 d, 14 d, and 21 d post‐injection. Ovaries from different groups were collected and fluorescent intensity was recorded using a small animal imaging system. Ovaries treated for 48 h were sectioned both longitudinally and transversely to evaluate the distribution and retention of EVs among varies groups. To confirm the chemotactic movement of nanomotors in LEVs‐Gel, we included a control group (Control + LEVs‐Gel) where LEVs‐Gel was administered to the ovaries of healthy mice. Observe the fluorescence distribution of the ovarian sections in each group using a fluorescence microscope.
In Vitro Maturation, Fertilization, and Embryo Culture of Oocytes
4.7
In vitro maturation (IVM) of oocytes was performed as described previously. Briefly, oocytes were isolated from mouse ovaries by puncturing follicles repeatedly and cultured in M2 medium for 2 h. Germinal vesicle breakdown (GVBD) was assessed, and oocytes were further cultured for 14–16 h to observe first polar body (PB1) extrusion, with maturation rates recorded. In vitro fertilization followed established protocols; C57 female mice were superovulated with gonadotropins and chorionic gonadotropin to induce ovulation. Sperm were collected from the cauda epididymis of male mice and capacitated in HTF medium. Cumulus‐oocyte complexes (COCs) were mixed with sperm in HTF droplets for fertilization. Post‐fertilization, zygotes were transferred to KSOM medium for continued culture. Embryo development was monitored, with two cell observed and counted at 24 h post‐fertilization and blastocysts formation assessed at 96 h.
Assessment of Reproductive Fertility Capacity
4.8
Natural mating experiments commenced 21 days post LEVs‐Gel transplantation. Twelve‐week‐old fertile C57 male mice were paired with treated female mice at a 1:2 ratio for 10 days, spanning two full estrous cycles to ensure adequate mating time. After the birth of the mouse pups, the reproductive fertility of the parent mice was evaluated by recording and comparing the fetal mortality, live birth count, sex ratio, and viability of each group.
Author Contributions
Yaoqin Mu: conceptualization, data curation, funding acquisition, writing – original draft, investigation, methodology, validation. Jiachen Wu: investigation, methodology, software, resources, validation. Mingwei Lv: methodology, software, data curation, resources, investigation. Xinyi Zhang: methodology, software, formal analysis. Yinhua Song: methodology, software, formal analysis. Jihui Ai: supervision. Ding Ma: funding acquisition, project administration, supervision. Kezhen Li: project administration, supervision, funding acquisition, writing – review and editing, validation.
Conflicts of Interest
Authors declare that they have no competing interests.
Supporting information
Additional experimental details, materials, and methods, including Supplementary Figures S1‐S6 as mentioned in the text (PDF).
Supporting Information: jev270249‐sup‐0002‐FigureS2.jpeg
Supporting Information: jev270249‐sup‐0003‐FigureS3.jpeg
Supporting Information: jev270249‐sup‐0004‐FigureS4.jpeg
Supporting Information: jev270249‐sup‐0005‐FigureS5.jpeg
Supporting Information: jev270249‐sup‐0006‐FigureS6.jpeg
Motion video of EVs (MP4).
Motion video of LEVs (MP4).
jev270249‐sup‐0009‐SuppMat.docx
The reference list from the paper itself. Each links out to its DOI / PubMed record.
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