MBs were fabricated using a lyophilization-based protocol. Hydrogenated soybean phospholipids (HSPC), hydrogenated egg yolk phospholipids (HEPC) and 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) were procured from Avite Pharmaceutical Technology Co., Ltd. (Shanghai, China). The lipid components were systematically formulated into six distinct groups. Lipid material (HSPC/HSPC/DSPC) (2.0 mg), Tween-80 (20.0 mg) (Solarbio, Beijing, China), and poloxamer 188 (200.0 mg) (Sigma‒Aldrich, MO, United States) were dissolved in 4.0 mL tert-butyl alcohol (XiLong Science Co., Ltd.). The solution was gradually cooled to complete solidification and subsequently stored at 4 °C overnight. Lyophilization was performed under reduced pressure (5 × 10⁻⁴ mBar) for 24 h using a three-stage protocol: primary drying at −48°C for 20 h, followed by a secondary drying phase with gradual temperature elevation to 5°C over 4 h. Finally, the powder obtained by freeze-drying was collected in a sterile container. For microbubbles preparation, aliquots of the powder were precisely weighed into vials. The gas exchange process was performed using vacuum-purge cycle displacement with either ambient air or NO gas (99.99%) to generate MBs or NO-MBs, respectively.

Microbubble suspensions were prepared by injecting 1 mL of phosphate-buffered saline (PBS, pH 7.4) into each vial containing 20.0 mg lyophilized powders. The six distinct formulations of MBs and NO-MBs were reconstituted in PBS and subjected to characterization. Morphological evaluation was conducted using an inverted optical microscope (Eclipse Ti2, Nikon), with representative images captured using NIS-Elements imaging software (v5.21). Quantitative analysis of particle size distribution and concentration was performed using a Multisizer 4e Coulter counter (Beckman Coulter Inc., Brea, CA, USA) with a 30 μm aperture tube. Measurements were conducted in triplicate for each formulation group under controlled temperature (25°C ± 0.5°C). Data acquisition and analysis were performed using the accompanying software (Multisizer 4e Software v3.01) with threshold settings optimized for microbubble detection.

The NO release profiles from MBs and NO-MBs were quantitatively evaluated using a dialysis method under sink conditions. Briefly, 1.0 mL of NO-MBs suspension (20 mg/mL, 2.23 × 10⁷ MBs/mL) was loaded into a dialysis bag (molecular weight cutoff: 3.5 kDa, Solarbio, Beijing, China) and immersed in 20 mL of phosphate-buffered saline (PBS, pH 7.4) as the receiving medium. Aliquots of the receiving buffer were collected at predetermined time intervals (5, 10, 20, 30, 40, 60, 80, 100, and 120 min) for NO quantification. The detection principle was based on the stoichiometric conversion of released NO to nitrite (NO₂ ⁻) via aerobic oxidation (4NO + O₂ + 2H₂O → 4NO₂ ⁻ + 4H⁺). The nitrite concentration in the receiving buffer was determined spectrophotometrically at 550 nm using a commercial NO assay kit (Solarbio, Beijing, China) according to the manufacturer’s protocol. All measurements were performed in triplicate to ensure data reproducibility.

Umbilical vein endothelial cells (HUVECs, ATCC^® CRL-1730™) were maintained in Dulbecco’s Modified Eagle Medium (DMEM, Gibco, USA) supplemented with 10% (v/v) fetal bovine serum (FBS, Gibco, USA) and 100 U/mL penicillin-streptomycin (Gibco, USA) at 37°C in a humidified 5% CO₂ incubator (Thermo Scientific, USA). Both murine and human cumulus granulosa cells were cultured in DMEM/F-12 medium (Gibco, USA). Test compounds—including MBs, NO-MBs, and sodium nitroprusside (SNP, Sigma-Aldrich, USA) as positive control—were administered to the culture medium at optimized concentrations and incubated with cells for 12 h under standard culture conditions. All cell culture procedures were performed under aseptic conditions in a Class II biological safety cabinet (Thermo Scientific, USA).

The cytotoxic effects of MBs and NO-MBs were evaluated using the MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] assay in both human cumulus granulosa cells and HUVECs (n = 6 replicates per group). Cells were seeded in 96-well culture plates at a density of 2 × 10⁴ cells/well in 200 μL complete growth medium and allowed to adhere for 12 h at 37°C in a 5% CO₂ atmosphere. Experimental groups were exposed to: MBs at logarithmic concentrations (1 × 10⁵, 1 × 10⁶, 1 × 10⁷ MBs/mL),NO-MBs at equivalent concentrations Serum-free medium as negative control,1% Triton X-100 as positive control. Following 12 h exposure, 20 μL MTT reagent was added per well and incubated for 4 h at 37°C. The formazan crystals were solubilized with 150 μL DMSO containing 0.01 M glycine buffer (pH 10.5). Absorbance was measured at 570 nm. Cell viability calculated as: Viability  % = A 570 , treated − A 570 , blank A 570 , control − A 570 , blank   × 100

The 3–4 weeks female mice were treated according to the guidelines (photoperiod of 12 h and free access to food and water). The mice were injected with PMSG (150 IU/kg). Intraperitoneally 48 h prior to dissection. Mice were anesthetized using chloral hydrate (Jurox Inc.) at 10 mg/kg body weight. The surgical site was disinfected using 70% ethanol, an incision was made on the linea alba, and the ovaries were retrieved, dissected and stored in prewarmed PBS solution. Later, the collected ovaries were sliced using a scalpel blade in M2 media. The released oocytes were then collected in 4-well plates containing 500 µL of M2 solution in each well. Only germinal vesicle oocytes that had three layers of unexpanded cumulus layers and balanced cytoplasm were collected. They were washed three times in M2 solution and then incubated in preprepared IVM media (M199 medium (Gibco, USA) supplemented with 0.1 mM sodium pyruvate, 75 mIU/mL FSH, 10% bovine serum albumin (Gibco, USA) and 1% penicillin-streptomycin and incubated) for maturation. Each COC was placed in a 10 µL droplet of IVM media, covered with mineral oil and incubated at 37°C and 5% CO₂ for predefined time periods; 18 h.

The NO levels in the oocytes were measured using the NO-sensitive fluorescent probe DAF-FM DA (Beyotime Biotechnology Inc.). Oocytes of each group were loaded with 5 µM DAF-FM DA at 37 °C for 30 min and then washed with PBS to remove the surface fluorescence. The intensity of the fluorescence in the whole oocyte was measured by laser scanning microscopy (LSM 800, Zeiss, Germany). The parameters used for image acquisition were similar for all examined oocytes. The experiments were independently repeated 3 times.

The ROS levels in IVM MII oocytes were measured using the ROS-sensitive fluorescent probe DCFH-DA (Beyotime Biotechnology Inc.). Oocytes of each group were loaded with 5 µM DCFH-DA at 37°C for 30 min and then washed with PBS to remove the surface fluorescence. The intensity of the fluorescence in the whole oocyte was measured by laser scanning microscopy (LSM 800, Zeiss, Germany). The parameters used for image acquisition were similar for all examined oocytes. The experiments were independently repeated 3 times.

The mitochondrial membrane potential (MMP) of the IVM MII oocytes was measured using JC-1 staining (Mitochon-Drial membrane potential assay kit, Abbkine Scientific Co., Wu Han, China). Briefly, oocytes from each group were exposed to JC-1 at 37°C for 20 min and then washed with PBS to remove surface fluorescence. The intensity of the fluorescence in the whole oocyte was measured by laser scanning microscopy (LSM 800, Zeiss, Germany). The parameters used for image acquisition were similar for all examined oocytes. The experiments were independently repeated 3 times.

The calcium levels in IVM MII oocytes were measured using the Ca²⁺-sensitive fluorescent probe Fluo-4 AM (Beyotime Biotechnology Inc.). Oocytes from each group were loaded with 5 µM Fluo-4 AM at 37°C for 30 min and then washed with PBS to remove surface fluorescence. The intensity of the fluorescence in the whole oocyte was measured by laser scanning microscopy (LSM 800, Zeiss, Germany). The parameters used for image acquisition were similar for all examined oocytes. The experiments were independently repeated 3 times.

Four-week-old female ICR mice were divided into four groups: MBs, NO-MBs, NO-MBs+L-NAME, and a control group (normal saline). MBs and NO-MBs were dissolved in normal saline to achieve a dose of 60 mg/kg, while the NO-MBs+L-NAME group received NO-MBs (60 mg/kg) combined with 10 mg/kg L-NAME. All groups received daily intraperitoneal injections for 7 days. Ovarian hyperstimulation was induced with PMSG (10 IU/mouse) followed by hCG (10 IU/mouse) 46–48 h later. All groups received hormone injections at synchronized time points. Immediately after hCG administration, females were cohoused with mature male mice. Vaginal plugs were checked 16–18 h post-mating, and plug-positive females were humanely euthanized simultaneously across all groups (n = 3 per group). Zygotes were collected from the ampulla of both fallopian tubes 20 h post-hCG injection (n = 40–60 zygotes per group), with all groups processed in parallel within a 30-min window to eliminate temporal confounding. After hyaluronidase-mediated granulosa cell removal, zygotes were cultured in pre-equilibrated embryo medium under oil at 37°C with 5% CO₂. Developmental stages (2-cell, 4-cell, morula, blastocyst) were assessed at standardized time points respectively. The whole procedure was repeated three times.

Protein quantification was performed using a bicinchoninic acid (BCA) assay. Equal amounts of protein lysates (20 μg per lane) were separated by 12% sodium dodecyl sulfate–polyacrylamide gel electrophoresis and transferred onto polyvinylidene difluoride membranes for Western blot analysis. After blocking with 5% bovine serum albumin for 2 h, the membranes were then incubated with anti-eNOS (1:500 dilution) (Affinity Biosciences), anti-iNOS (1:200 dilution) (Affinity Biosciences), anti-ERK 1/2 (1:1,000 dilution), and anti-p-ERK1/2 (1:1,000 dilution) antibodies as primary antibodies at 4°C overnight. The membranes were washed before incubation with HRP-conjugated goat anti-IgG (H+L) secondary antibody (1:10,000 dilution) for 1 h at room temperature. The relative band density was calculated using ImageJ software, and an antibody against GAPDH was used as an internal control.

The data are presented as the mean ± standard deviation (SD) and were analyzed using GraphPad Prism 8.0 (GraphPad Software, USA). Significant differences between two groups were determined by an unpaired Student’s t-test, while multiple comparisons were assessed by one-way ANOVA followed by Tukey’s post hoc test. A p-value <0.05 was considered statistically significant.

Six lipid formulations were systematically evaluated for microbubble stability and gas encapsulation potential. Single-component systems (Group 1: HSPC; Group 2: HEPC; Group 3: DSPC) and combinatorial formulations (Group 4: HSPC+HEPC; Group 5: HEPC+DSPC; Group 6: HSPC+DSPC) were prepared via lyophilization (Figures 2A, B) and characterized.

All formulations exhibited homogeneous curd-like suspensions, with visually distinct turbidity levels observed across the groups (Figure 2C). Particle size distribution and concentration analysis revealed that combination groups exhibited higher particle numbers compared to single-component groups (Figures 2B, D). Among these, Group 6 (HSPC + DSPC) demonstrated the highest particle concentration (2.23 × 10⁷ MBs/mL). Optical microscopy analysis revealed distinct variations in particle density across the groups, with Group 6 demonstrating both higher particle concentration and superior size uniformity compared to other formulations (Figure 2E). To assess stability, the particle numbers of 20 mg/mL microbubble solutions were measured after 24 h at room temperature. Group 6 maintained the highest particle number [(67.00 ± 14.38) ×10⁵ MBs/mL, p < 0.05 vs. all groups (Figure 2F)], indicating superior stability. In summary, the combination of HSPC and DSPC was used as the lipid material for microbubbles.

Based on preliminary screening of carrier materials for NO gas delivery (Section 3.1), NO-loaded microbubbles (NO-MBs) were constructed and characterized for stability, biological safety, and functional efficacy. The NO loading methodology is illustrated in Figure 3A. Both blank microbubbles (MBs) and NO-MBs exhibited homogeneous colloidal suspensions (100 mg/mL in PBS) with equivalent macroscopic phase behavior (Figure 3B). Microscopic examination revealed polydisperse spherical particles with no evidence of aggregation or structural deformation post-NO loading (Figure 3C).

Cytotoxicity assessment demonstrated excellent biocompatibility of NO-MBs across a concentration range of 0–5 mg/mL (Figure 3D). Notably, low-dose exposure (≤0.5 mg/mL) exhibited a mild proliferative effect on HUVECs (viability = 118.38 ± 55.63% vs. control, p < 0.01), potentially attributable to NO-mediated endothelial cell activation.

In vitro release profiles of NO-MBs were quantified using chemiluminescence detection (Figure 3E). Blank MBs showed baseline NO levels, confirming encapsulation specificity. NO-MBs exhibited a biphasic release profile: an initial burst phase (0–60 min) achieving maximal NO concentration (21.70 ± 0.17 μM), followed by sustained release maintaining therapeutic levels (16.6 ± 1.13 μM) over 24 h. These results demonstrated that NO-MBs enable stable, sustained, and biologically safe NO delivery.

To evaluate the reproductive potential of NO-MBs, GV-stage oocytes were isolated from 3-4-week-old ICR mice following superovulation (10 IU PMSG, 46–48 h) and cultured in IVM medium supplemented with NO-MBs or SNP. Oocyte retrieval was performed via ovarian dissection post-cervical dislocation (Figure 4A). And as shown in Figure 4B, morphological analysis of oocytes at different developmental stages revealed distinct characteristics.

Osmolarity measurements provided critical insights into culture conditions: While NO-MBs supplementation maintained physiological osmolarity (306.00 ± 2.00 mOsm/kg vs. control 308.67 ± 0.58 mOsm/kg, P = 0.21), SNP treatment significantly elevated medium osmolarity (315.00 ± 1.00 mOsm/kg, P < 0.001 vs. control) (Figure 4C), potentially contributing to its detrimental effects on oocyte maturation. All of the GV oocytes were collected and divided into three groups (Control, NO-MBs and SNP) in the subsequent IVM experiments. As shown in Figure 4D, morphological evaluation of oocytes after IVM culture revealed superior cytoplasmic quality in the NO-MBs-treated group compared to other groups. Quantitative analysis demonstrated that NO-MBs supplementation significantly improved the maturation rate of immature oocytes (72.00%) compared to the control group (57.77%, P = 0.003) (Figure 4E). In contrast, the NO donor SNP treatment resulted in a reduced maturation rate relative to the control group.

Microscopic observation revealed that oocytes in the NO-MBs group exhibited superior morphological characteristics compared to the other groups. To further evaluate the quality and developmental potential of these oocytes, we analyzed several key biomarkers associated with oocyte competence.

Intracellular reactive oxygen species (ROS), critical for spindle formation, chromosome alignment, and oocyte maturation (Jiao et al., 2021), were significantly reduced by NO-MBs treatment compared to controls (n = 7; P < 0.05). In contrast, SNP-treated oocytes (n = 6) exhibited elevated ROS levels versus both control (P < 0.01) and NO-MBs groups (P < 0.001) (Figures 5A, D).

Calcium (Ca²⁺) signaling, essential for meiotic resumption and germinal vesicle breakdown (Deguchi et al., 2015), was enhanced by NO-MBs (n = 8; P < 0.01 vs. control), while SNP treatment (n = 5) suppressed Ca²⁺ levels (P < 0.05 vs. control) (Figures 5B, E). This suggests NO-MBs promote Ca²⁺ oscillations, facilitating meiotic resumption and cortical granule exocytosis—key mechanisms for preventing polyspermy.

Mitochondrial function, assessed via membrane potential (MMP) using JC-1 fluorescence, revealed significantly increased MMP in NO-MBs-treated oocytes (Figures 5C, F). JC-1 aggregates (red fluorescence, ΔΨm >140 mV) increased relative to monomers (green fluorescence, ΔΨm <100 mV), indicating enhanced mitochondrial polarization critical for energy production and developmental competence (Kirillova et al., 2021).

To confirm the effect of NO-MBs on intracellular NO levels of oocytes, we measured the concentration of NO in the IVM-MII oocytes of each group. Quantitative analysis revealed that both NO-MBs and SNP treatments significantly enhanced NO generation compared to controls (n = 10). Specifically, mean fluorescence intensity increased in the NO-MBs group (n = 7) and SNP group (n = 6) (P < 0.001; Figures 6A, B), confirming NO-MBs as an effective NO donor in oocytes.

To determine whether NO-MBs promote maturation through nitric oxide synthase (NOS)-mediated NO synthesis, we employed L-NAME, a specific NOS inhibitor. Immature oocytes were divided into six experimental groups: Control, MBs, NO-MBs, NO-MBs + L-NAME, SNP, SNP + L-NAME (Figure 6C). Maturation rates demonstrated a NOS-dependent mechanism for NO-MBs: NO-MBs significantly enhanced maturation (86.6% vs. 75% control; P < 0.01); L-NAME abolished this effect (66.7% vs. 86.6%; P < 0.01), even reducing maturation below control levels. In contrast, L-NAME paradoxically improved maturation in SNP-treated oocytes (78.5% vs. 66.7%; P < 0.05; Figure 6D). These findings reveal the dual role of NO in oocyte development, where endogenous (NOS-mediated) and exogenous NO sources exert distinct biological effects.

Western blot analysis of ERK1/2 phosphorylation status provided mechanistic insights: NO-MBs uniquely enhanced p-ERK1/2 expression (vs. control; P < 0.01). Total ERK1/2 levels remained stable across groups (NS). Phosphorylation ratio (p-ERK/ERK) was highest in the NO-MBs group (Figures 6E, F). This selective ERK activation suggests that NO-MBs promote maturation through both NO release and downstream MAPK signaling modulation. In summary, NO-MBs enhance oocyte maturation through: sustained NO release, NOS-dependent endogenous NO synthesis, selective activation of ERK signaling pathway. These findings establish NO-MBs as a multifaceted modulator of oocyte quality, offering potential applications in assisted reproductive technologies.

To comprehensively evaluate the safety and efficacy of NO-MBs on embryonic development, we conducted a systematic in vivo study in female mice. Following 1 week of continuous intraperitoneal administration (5 mg/kg/day), oocytes were retrieved and subjected to in vitro fertilization (IVF) and embryo culture. NO-MBs treatment significantly improved developmental parameters compared to controls: fertilization rate: 86.32% (NO-MBs) vs. 70.04% (Control) (**P < 0.01). Blastocyst formation rate: 94.42% vs. 73.48% (***P < 0.001) (Figures 7A, B).

TUNEL/DAPI double staining was used to detect the apoptosis rate of blastocyst cells. The results showed that the apoptosis rate of blastocyst cells in the NO-MBs (71.0%) intraperitoneal injection group was significantly lower than that in the control group (86.5%) and MBs injection group (76.2%) (Figures 8A, B).