US-triggered on-demand NO-releasing biomimetic nanoparticle to Remodel endothelial microenvironment for enhancing atherosclerosis-specific gas therapy

Abstract

Atherosclerosis (AS) is a chronic inflammatory disease driven by endothelial dysfunction, vascular smooth muscle cell proliferation, and insufficient resolution of inflammation. Nitric oxide (NO) plays a crucial role in vascular homeostasis by promoting endothelial cell proliferation, maintaining endothelial integrity, suppressing smooth muscle cell hyperplasia, and exerting potent anti-inflammatory effects. However, clinical application of NO is hindered by its short half-life, lack of targeting, and uncontrolled release. Here, we developed the biomimetic nanoparticles (B-NPs@MM) for targeted and controllable NO delivery by encapsulating the ultrasound (US)-responsive NO donor BNN6 into poly(lactic-co-glycolic acid) (PLGA) nanospheres followed by coating with macrophage-derived membranes. These biomimetic particles mimic natural macrophages to actively target inflamed atherosclerotic plaques and evade immune clearance. Upon localized US exposure, the system triggers rapid and on-demand NO release at the lesion site with spatiotemporal precision. In vitro and in vivo evaluations demonstrate effective NO delivery, enhanced endothelial repair, reduced inflammation, and inhibition of neointimal hyperplasia. This work presents a smart, remotely controlled NO nanotherapy platform based on artificial immune cell design, offering a promising strategy for precision treatment of AS.

Graphical abstract

Highlights

• •Ultrasound-triggered NO release enables spatiotemporal control.
• •B-NPs@MM promotes endothelial repair and suppresses inflammation.
• •Achieves early-stage anti-atherosclerotic effects in vivo.

1. Introduction

Atherosclerosis (AS) is a chronic inflammatory vascular disease characterized by lipid accumulation, endothelial dysfunction, immune cell infiltration, and smooth muscle cell proliferation [1,2]. As the leading underlying cause of myocardial infarction, stroke, and peripheral artery disease, AS remains a major contributor to cardiovascular mortality worldwide [3]. While current clinical interventions such as statins, anti-inflammatory agents, and revascularization procedures alleviate symptoms and slow progression, they are often insufficient in reversing plaque formation or restoring vascular homeostasis [4,5]. Therefore, the development of targeted, mechanism-informed therapeutic strategies remains an urgent need.

Recent studies have emphasized the therapeutic potential of bioactive systems, particularly those inspired by natural products, in treating cardiovascular disorders by modulating inflammation, oxidative stress, and vascular remodeling [2]. Among these strategies, nanotechnology-based platforms offer significant advantages in delivering therapeutic agents with spatiotemporal precision [6]. Nitric oxide (NO), a gaseous signaling molecule endogenously produced by endothelial cells, plays a critical role in maintaining vascular homeostasis and preventing atherosclerotic progression. Therefore, developing nanoplatforms capable of targeted and controllable NO delivery presents a promising avenue for cardiovascular therapy [7,8]. It promotes endothelial cell proliferation and migration, enhances endothelial barrier function, inhibits VSMC proliferation, suppresses oxidative stress, and modulates immune responses by shifting macrophage polarization from pro-inflammatory M1 to anti-inflammatory M2 phenotypes [[9], [10], [11]]. Moreover, NO reduces platelet adhesion and aggregation, thereby preventing thrombosis. These diverse effects underscore NO's therapeutic potential in AS management [12]. However, the clinical application of NO is constrained by its extremely short half-life and high diffusibility, which hinder its localization and concentration control [13,14]. Notably, NO exhibits a dose-dependent duality: at physiological levels, it supports cellular repair and angiogenesis, whereas excessive concentrations can induce cytotoxicity and inflammation. Therefore, spatially and temporally controlled delivery of NO is essential for maximizing its therapeutic efficacy while minimizing systemic side effects [15,16].

To address these challenges, stimulus-responsive delivery systems have gained attention for enabling precise NO release [14,17]. Among various external stimuli, ultrasound (US) stands out for its noninvasive nature, deep tissue penetration, and real-time controllability [18,19]. US-triggered NO donors, such as N,N′-di-sec-butyl-N,N′-dinitroso-1,4-phenylenediamine (BNN6), can decompose upon US exposure to release NO in a controlled manner [20]. This mechanism allows for remote, site-specific activation of NO delivery, making it particularly advantageous for the treatment of spatially localized vascular lesions such as atherosclerotic plaques. We have previously demonstrated a similar ultrasound-responsive NO delivery strategy in the context of myocardial infarction, highlighting its potential in localized cardiovascular therapy [21,22].

In the context of AS, macrophages play a pivotal role in both disease progression and resolution [23]. They are among the important immune cells recruited to the intimal layer and contribute to foam cell formation, oxidative stress, and pro-inflammatory cytokine secretion [24]. However, macrophages also possess natural plaque-homing behavior mediated by surface adhesion molecules and chemokine receptors [25]. Harnessing this property, biomimetic nanoparticles coated with macrophage membranes can achieve active targeting of inflammatory vascular sites [26]. Additionally, macrophage-derived membranes retain immune-modulatory proteins such as CD47, which signal “self” to phagocytes, thereby reducing mononuclear phagocyte system clearance and prolonging systemic circulation [27]. These features make macrophage membrane camouflage a compelling strategy for delivering therapeutic agents to atherosclerotic plaques with high specificity and minimal immunogenicity [28].

To realize this concept, we developed a biomimetic, US-responsive nanoparticle termed artificial macrophage biomimetic nanoparticle (B-NPs@MM). As shown in Fig. 1A, the system consists of BNN6-loaded poly(lactic-co-glycolic acid) (PLGA) nanoparticles cloaked with natural macrophage membranes. The biomimetic nanoparticle combine immune-evasive and inflammation-targeting capabilities with US-triggered, on-demand NO release. Upon activation, localized NO generation exerts a multifaceted therapeutic effect by promoting endothelial repair, inhibiting vascular smooth muscle cell proliferation, and modulating the local immune response (Fig. 1B). This strategy offers a promising approach for spatiotemporally precise, biomimetic gas therapy in AS.

Fig. 1.

Schematic illustration of B-NPs@MM construction and their therapeutic mechanism for atherosclerosis. (A) Fabrication of biomimetic nanoparticle B-NPs@MM. The system consists of PLGA nanoparticles encapsulating the US-responsive NO donor BNN6 (B-NPs), which are subsequently cloaked with macrophage-derived membranes (MM). (B) Therapeutic mechanism of B-NPs@MM in atherosclerosis. After homing to inflamed atherosclerotic plaques, US irradiation induces on-demand NO release from the nanocarrier core. The released NO promotes endothelial cell (EC) proliferation and repair, inhibits smooth muscle cell (SMC) hyperproliferation, and mitigating vascular inflammation and stabilizing atherosclerotic lesions.

2. Results and discussion

2.1. Synthesis and characterization of B-NPs@MM

The chemical structure of the synthesized BNN6 was confirmed by electrospray ionization mass spectrometry (Fig. S1), which exhibited a prominent molecular ion peak at m/z = 279, consistent with its theoretical molecular weight, thereby verifying the successful synthesis of BNN6 [29]. As illustrated in the reaction scheme (Fig. S2), BNN6 undergoes homolytic cleavage of the N–NO bond under US irradiation, releasing two molecules of NO per BNN6 molecule. This US-triggered NO release mechanism ensures controllability and minimal background leakage, offering spatiotemporal precision for therapeutic delivery [20].

B-NPs@MM, a macrophage membrane-coated NO delivery nanoparticle, was constructed through BNN6 encapsulation into PLGA nanoparticles followed by extrusion with macrophage membranes, as shown in Fig. 2A. The drug-loading capacity (DL%) of BNN6 in B-NPs was 12.4 %. The encapsulation efficiency (EE%) of BNN6 was 74.2 %, these results demonstrate satisfactory drug loading and high encapsulation efficiency, supporting the successful incorporation of BNN6 into our nanoplatform. This system is capable of US-triggered NO release. Transmission electron microscopy (TEM) revealed that both B-NPs and B-NPs@MM exhibited uniform, spherical morphology with smooth surfaces and average diameters around 200 nm. Notably, B-NPs@MM displayed a distinct core–shell structure, confirming the successful coating of macrophage membranes onto the nanoparticle surface (Fig. 2B). Dynamic light scattering (DLS) analysis showed that B-NPs@MM exhibited an average hydrodynamic diameter of 232.4 ± 3.5 nm with a low polydispersity index (PDI = 0.219), indicating uniform particle distribution and good colloidal stability (Fig. 2C). The average diameters of PLGA nanoparticles and B-NPs were 228.9 ± 6.3 nm and 221.1 ± 6.2 nm, respectively, suggesting that membrane coating led to a slight increase in particle size. This increase is consistent with the presence of an outer membrane layer and corroborates the TEM observations of a core–shell morphology. Zeta potential measurements further supported successful membrane fusion. The surface charge of PLGA nanoparticles was −22.0 ± 3.3 mV, which increased to −18.3 ± 1.68 mV upon BNN6 encapsulation (B-NPs), and further increased to −9.6 ± 4.1 mV after membrane coating (B-NPs@MM) (Fig. 2D). The zeta potential of the extracted macrophage membrane vesicles was −10.4 ± 1.7 mV, closely matching that of B-NPs@MM. These results suggest that the negatively charged PLGA core was effectively shielded by the relatively less negatively charged macrophage membrane, a phenomenon consistent with previous studies on membrane-camouflaged nanocarriers [30]. The observed shift in surface potential, together with the unchanged size distribution, further confirms the successful fabrication of B-NPs@MM.

Fig. 2.

Characterization of B-NPs@MM and their US-responsive NO release properties. (A) Schematic illustration of the preparation of B-NPs@MM, including BNN6-loaded PLGA core formation, macrophage membrane coating, and US-responsive NO release. (B) TEM images of B-NPs and B-NPs@MM. (C–D) DLS and zeta potential measurements of NPs, B-NPs, B-NPs@MM, and RAW264.7 membranes. (E) FTIR spectra of BNN6, NPs, and B-NPs. (F–G) TGA and DSC curves of BNN6, NPs, and B-NPs. (H) Western blot analysis of integrin α4, integrinβ1, and CD47. (I) US-triggered NO release profiles of BNN6, B-NPs, and B-NPs@MM over time.

Fourier-transform infrared (FTIR) spectroscopy was conducted to confirm the successful incorporation of BNN6 into the nanoparticle system. As shown in Fig. 2E, the characteristic absorption peak of BNN6 at 1512 cm⁻¹ and 1012 cm⁻¹, corresponding to the –N–N=O stretching vibration and aromatic C–N bonds, was clearly observed in both the free BNN6 and the B-NPs spectrum. This peak remained detectable, though slightly attenuated, in the B-NPs@MM group, indicating that BNN6 was successfully encapsulated and retained its structural integrity during nanoparticle fabrication and membrane coating. The reduction in peak intensity is attributed to the relatively low BNN6 loading content and the presence of PLGA and membrane components, which may partially mask BNN6 signals. Nonetheless, the appearance of the 1511 cm⁻¹ and 1012 cm⁻¹ peak across BNN6, B-NPs, and B-NPs@MM confirms that the characteristic chemical features of BNN6 were preserved throughout the preparation process. These findings provide strong evidence for the successful loading of BNN6 into the PLGA core and further suggest that the macrophage membrane coating does not interfere with the molecular identity of the encapsulated compound.

To further validate the chemical stability and encapsulation state of BNN6 within the PLGA matrix and upon macrophage membrane coating, we conducted Raman spectroscopy on B-NPs@MM, B-NPs, PLGA, and free BNN6. As shown in Fig. S3, the characteristic BNN6 peaks at ∼1535 cm⁻¹ (N-NO symmetric stretch) and ∼1600 cm⁻¹ (C=C stretching) remained clearly identifiable in both B-NPs and B-NPs@MM, with negligible shifts (<2 cm⁻¹) and minimal peak area variation compared to free BNN6, indicating that the molecular structure of BNN6 was well preserved during the encapsulation and coating processes. In addition, the C=O stretching peak of PLGA shifted slightly from 1748 cm⁻¹ to 1745 cm⁻¹ upon BNN6 loading, suggesting weak hydrogen bonding interactions without covalent modification. The exclusive appearance of a pronounced Amide I band at ∼1644 cm⁻¹ in B-NPs@MM further confirmed the successful membrane coating. Notably, the slight broadening of this band, in the absence of significant peak shifts, implies minimal perturbation to membrane protein structure, suggesting that BNN6 encapsulation did not induce direct chemical interactions with membrane components. These Raman results, together with FTIR and encapsulation efficiency data (DL = 12.4 %, EE = 74.2 %), comprehensively support the successful and stable integration of BNN6 into the nanoparticle system, while maintaining chemical integrity and biocompatible surface features.

Thermogravimetric analysis (TGA) and differential scanning calorimetry (DSC) were employed to assess the thermal stability and composition of B-NPs. As shown in Fig. 2F, BNN6 exhibited a major weight loss beginning at ∼150 °C due to its thermal decomposition. In contrast, blank NPs showed thermal stability up to ∼300 °C. B-NPs displayed a two-step degradation profile, with an early weight loss between 150 and 250 °C attributable to encapsulated BNN6, confirming successful drug loading. The corresponding DSC curves (Fig. 2G) further support these findings. BNN6 showed a sharp endothermic peak around 75 °C and a broad exothermic event near 200 °C, indicating melting and decomposition. Blank NPs exhibited a dominant thermal transition around 310 °C, corresponding to PLGA matrix degradation. Notably, B-NPs displayed both BNN6-related and polymer-related transitions, with a shift in peak positions, suggesting BNN6 was molecularly dispersed within the polymer and its thermal behavior was altered upon encapsulation. Together, these results confirm that BNN6 was effectively integrated into the nanoparticle matrix without compromising the thermal stability of the carrier system.

SDS-PAGE (Fig. S4) and Western blotting (Fig. 2H) demonstrated that B-NPs@MM preserved key macrophage membrane proteins such as integrin β, CD11b and CD47. These proteins are essential for macrophage recognition, adhesion to inflammatory endothelium, and phagocytosis evasion [31,32]. The retention of functional proteins indicates the structural and biological integrity of the membrane coating process.

Collectively, these results confirm the successful construction of B-NPs@MM with stable morphology, effective BNN6 loading, preserved membrane functionality, and US-triggered NO release. Compared with other nanoparticle-based NO delivery systems, the integration of macrophage-derived membranes offers distinct advantages in targeting inflamed vasculature and evading clearance by the mononuclear phagocyte system [33,34]. This design holds significant promise for non-invasive, precise, and controllable NO therapy, particularly in the context of AS or other inflammatory vascular diseases.

2.2. US-triggered NO release from B-NPs@MM

The US-responsive NO release capacity of B-NPs@MM was evaluated using the Griess assay. Upon US irradiation (1.0 W/cm², 50 % duty cycle, 1 MHz), B-NPs@MM exhibited a time-dependent NO release profile, similar to free BNN6 and B-NPs, while negligible NO release was observed without US stimulation.

As shown in Fig. 2I, the concentration of NO released from free BNN6, B-NPs, and B-NPs@MM (all at BNN6-equivalent concentration of 100 μM) increased progressively with the extension of US irradiation time. After 60 min of cumulative irradiation, all formulations exhibited significant NO release, with BNN6 reaching a maximum concentration of approximately 65 μM. The slight underestimation of final NO levels may be attributed to partial conversion of NO to NO₂⁻ during detection and volatilization losses during sonication [21]. To assess whether macrophage membrane coating affects NO release, we compared B-NPs and B-NPs@MM under the same US stimulation. As shown in Fig. 2I, both showed rapid NO release, with B-NPs@MM exhibiting only a slight initial delay. This minor difference may stem from the membrane acting as a transient barrier, but US likely enhanced membrane permeability, enabling efficient NO diffusion. Thus, membrane coating had minimal impact on release kinetics, confirming B-NPs@MM as a viable NO delivery system.

Compared to free BNN6, both B-NPs and B-NPs@MM released NO at a slower rate during the initial 20 min of US exposure. This delay is likely due to the encapsulation of BNN6 within the PLGA matrix, which acts as a diffusion barrier and modulates the release kinetics. Notably, the NO release profile of B-NPs@MM was comparable to that of uncoated B-NPs, suggesting that the macrophage membrane coating did not significantly hinder US-responsive BNN6 decomposition.

These results demonstrate that B-NPs@MM enables dose-controllable, externally regulated NO release under US stimulation. This controllability is advantageous for cardiovascular applications, where excessive NO can lead to cytotoxic effects, and precise NO dosing is crucial for maintaining therapeutic efficacy and safety.

2.3. In vitro biosafety

To evaluate blood compatibility, a standard hemolysis assay was performed using red blood cells incubated with B-NPs and B-NPs@MM at varying concentrations (0.1, 1, and 5 mg/mL). As shown in Fig. S5, the nanoparticle suspensions remained clear with no visible red discoloration, while the Triton X-100 group showed intense hemoglobin release, serving as the positive control. Quantitative analysis revealed that hemolysis rates in both B-NPs and B-NPs@MM groups were consistently below 5 %, comparable to the PBS-treated negative control, indicating good erythrocyte tolerance and low hemolytic potential. These results indicate excellent red blood cell compatibility in accordance with accepted biomaterials safety standards [35].

The cytotoxicity of both formulations was further investigated in three representative cell types: HUVECs, VSMCs and RAW264.7 macrophages. As depicted in Fig. S6, exposure to a range of nanoparticle concentrations (0.1–10 mg/mL, normalized to BNN6 content) for 24 h did not induce significant loss of viability in any cell line. All groups maintained cell viability above 90 %, and no significant differences were observed between membrane-coated and uncoated formulations.

Importantly, the macrophage membrane camouflage in B-NPs@MM did not introduce additional cytotoxicity, which may be attributed to its biological origin and immunologically inert surface profile [36]. Membrane-cloaked nanocarriers have previously been shown to enhance biocompatibility while enabling immune evasion, making them attractive for systemic administration [37]. The favorable biocompatibility observed here supports the feasibility of B-NPs@MM for in vivo applications, including vascular-targeted drug delivery and NO-based AS therapy.

2.4. Assessment of the targeting capability of B-NPs@MM In vitro and In vivo

To evaluate the inflammation-targeting ability of B-NPs@MM, we examined their binding behavior on H₂O₂-stimulated endothelial cells. As shown in Fig. 3A, minimal DiI fluorescence was observed in both B-NPs and B-NPs@MM groups under normal (H₂O₂−) conditions. However, in H₂O₂-treated HUVECs, B-NPs@MM exhibited markedly enhanced surface adhesion compared to uncoated B-NPs, as evidenced by stronger fluorescence signals. Quantitative image analysis confirmed significantly greater adhesion of B-NPs@MM to inflamed HUVECs (P < 0.001), while no difference was found in the unstimulated control group (Fig. 3B). This indicates that the macrophage membrane confers inflammation-responsiveness and preferential adhesion to activated endothelial cells. To further verify the targeting performance of B-NPs@MM in endothelial cells, flow cytometry was conducted in HUVECs. As shown in Fig. S7, the DiD-positive cell population for B-NPs@MM (51.0 %) was markedly higher than that of uncoated B-NPs (30.4 %), demonstrating enhanced cellular banding efficiency after macrophage membrane camouflage.

Fig. 3.

In vitro and in vivo targeting and adhesion performance of B-NPs@MM. (A) Fluorescence microscopy images showing the binding of DiI-labeled B-NPs and B-NPs@MM to HUVECs under normal (H₂O₂−) and inflammatory (H₂O₂+) conditions. (B) Quantitative analysis of nanoparticle adhesion to HUVECs based on mean fluorescence intensity. (C) Representative fluorescence images of VSMCs incubated with B-NPs or B-NPs@MM under H₂O₂ stimulation. (D) Quantification of nanoparticle adhesion to VSMCs. (E) Flow cytometry analysis of RAW264.7 macrophages after incubation with DiI-labeled B-NPs or B-NPs@MM, showing reduced nonspecific association of B-NPs@MM. (F) Ex vivo fluorescence imaging of cardiovascular tissue 24 h after intravenous injection of DiR-labeled nanoparticles. (G) Quantification of organ fluorescence intensity.

A similar adhesion pattern was observed in H₂O₂-treated vascular smooth muscle cells (VSMCs), where B-NPs@MM demonstrated stronger binding than B-NPs under inflammatory conditions (Fig. 3C and D). This adhesion enhancement is likely due to the presence of membrane-associated adhesion molecules, such as integrin α4 and integrin β1, retained on the macrophage membrane surface [31,33].

To investigate immune evasion, RAW264.7 macrophages were incubated with DiI-labeled B-NPs and B-NPs@MM, and associated fluorescence was analyzed by flow cytometry. As shown in Fig. 3E, B-NPs@MM showed significantly lower fluorescence compared to B-NPs (16.7 % vs. 50.7 %), suggesting reduced nonspecific binding or phagocytic interaction, likely due to the presence of CD47, which provides a “self-recognition” signal and inhibits clearance by macrophages [38].

To further assess the in vivo vascular targeting ability, DiR-labeled nanoparticles were intravenously injected into ApoE^−/− mice, and near-infrared fluorescence imaging was performed 24 h post-injection (Fig. 3F). Compared to the B-NPs group, the B-NPs@MM group showed markedly stronger fluorescence signals along the aortic region, indicating enhanced vascular accumulation. Quantitative analysis (Fig. 3G) revealed a significantly higher average radiant efficiency in the B-NPs@MM group than in the B-NPs group (P < 0.05), confirming improved targeting efficiency. The enhanced vascular localization is attributable to the macrophage membrane coating, which facilitates lesion-specific adhesion via recognition of inflammatory endothelium and chemokine gradients [33]. This biomimetic feature allows B-NPs@MM to preferentially accumulate at atherosclerotic sites, thereby laying the foundation for localized NO release and lesion-specific therapy.

2.5. In vitro evaluation of endothelial regeneration and VSMCs suppression

Restoring endothelial integrity at the early stage of AS is crucial for maintaining vascular homeostasis and preventing plaque initiation [39]. Endothelial cell proliferation and migration are essential processes in repairing vascular injury, maintaining barrier function, and reducing lipid deposition and inflammatory infiltration [40]. Endothelial dysfunction, characterized by impaired regenerative capacity and increased permeability, is a hallmark of early AS progression [41].

To investigate the regenerative potential of B-NPs@MM, a series of in vitro assays were performed on HUVECs under oxidative stress conditions. In the wound healing assay (Fig. 4A and B), cells treated with B-NPs@MM + US showed a significantly accelerated migration rate compared to the control group (P < 0.001), while no significant difference was observed between the B-NPs@MM and control groups. This highlights the essential role of US-triggered NO release in enhancing endothelial motility. Similarly, the Transwell migration assay (Fig. 4C and D) demonstrated that B-NPs@MM + US significantly increased the number of migrated cells compared to both the control and B-NPs@MM groups (P < 0.001), further confirming the efficacy of NO delivery in promoting endothelial cell migration. Again, the B-NPs@MM group did not show a significant improvement over the control, suggesting that passive targeting alone was insufficient under oxidative conditions. In the tube formation assay (Fig. 4E–G), B-NPs@MM + US substantially enhanced the formation of vascular-like networks, as evidenced by increased junction numbers and total branch length (P < 0.05 vs. control), whereas B-NPs@MM had minimal effect. These findings underscore the importance of controlled NO release in stimulating angiogenesis. Cell proliferation, assessed by the EdU assay (Fig. 4H and I), was also significantly improved in the B-NPs@MM + US group compared to the control and B-NPs@MM groups (P < 0.001), indicating that US-mediated NO delivery effectively promoted endothelial proliferation.

Fig. 4.

Evaluation of HUVEC migration, proliferation, and angiogenesis after treatment. (A) Representative images of scratch wound healing assay at 0 h and 24 h post-treatment. (B) Quantification of wound closure rate based on migration area (%). (C) Representative images of transwell migration assay stained with crystal violet. (D) Quantitative absorbance analysis of migrated cells. (E) Representative images of tube formation on Matrigel after treatment. (F) Quantification of tube junction numbers. (G) Quantification of total branch length. (H) Representative EdU fluorescence images showing proliferative HUVECs. (I) Quantification of EdU⁺ cells expressed as percentage of total cells.

In parallel, we investigated the effect of NO on VSMCs behavior. As shown in Fig. 5A and B, the EdU assay revealed that B-NPs@MM + US significantly reduced the proliferation of VSMCs compared with control and B-NPs@MM groups (P < 0.001), aligning with NO's known inhibitory effects on SMC hyperplasia [42]. These data confirmed the anti-proliferative effect of B-NPs@MM-mediated NO release on activated VSMCs. It has been reported that NO can trigger p53/p21 pathway activation, leading to cell cycle arrest and reduced proliferation in VSMCs [43]. The observed inhibition effect in our study could be partially attributed to this mechanism. This supports the therapeutic potential of B-NPs@MM + US in limiting neointimal thickening and plaque progression.

Fig. 5.

Assessment of anti-proliferative effects on VSMCs and anti-inflammatory effects on H₂O₂-stimulated HUVECs.(A) Representative fluorescence images of EdU staining in VSMCs after different treatments. (B) Quantification of EdU⁺ VSMCs expressed as a percentage of total cells. (C) Representative fluorescence images of intracellular ROS levels in H₂O₂-pretreated HUVECs using DCFH-DA staining. (D) Quantification of ROS fluorescence intensity expressed in arbitrary units (a.u.).

Furthermore, ROS staining in H₂O₂-pretreated HUVECs (Fig. 5C and D) showed that B-NPs@MM + US significantly suppressed intracellular ROS accumulation relative to control and B-NPs@MM groups (P < 0.001), reflecting the antioxidant and anti-inflammatory effects of NO, which helps maintain endothelial homeostasis under inflammatory insult [44].

To differentiate the contribution of exogenous NO released from B-NPs@MM from endogenous NO production, we conducted DAF-FM DA fluorescence imaging and cGMP ELISA assays in both HUVECs and RAW264.7 cells. As shown in Figs. S8 and S9, H₂O₂ treatment significantly reduced intracellular NO levels in HUVECs, likely due to oxidative suppression of eNOS activity, while LPS stimulation transiently increased NO levels in RAW264.7 cells via iNOS induction. However, in both cases, intracellular cGMP levels remained unchanged, indicating that endogenous NO alone was insufficient to elicit a downstream biological response. In contrast, B-NPs@MM significantly increased both NO fluorescence intensity and cGMP levels following US activation, even in the presence of NOS inhibitors (L-NAME for HUVECs, 1400W for RAW cells), suggesting that the observed signaling effects were predominantly derived from exogenous NO. Notably, B-NPs@MM + US restored H₂O₂-impaired cGMP production in HUVECs, confirming the ability of the released NO to activate the sGC–cGMP axis under oxidative conditions. These findings support the functional bioactivity of exogenous NO delivered by B-NPs@MM and are consistent with previous reports using graphene-based NO delivery platforms, which also demonstrated cGMP-mediated endothelial protection and smooth muscle inhibition via intracellular NO release [45]. Together, these results validate the biological relevance of NO release from our system and underscore its therapeutic potential for vascular protection under pathological conditions.

Together, these findings demonstrate that B-NPs@MM + US significantly enhances endothelial regeneration, suppresses VSMCs proliferation, and alleviates oxidative stress, offering a comprehensive therapeutic approach for early-stage AS. The ability to spatiotemporally control NO release via US further ensures safety and precision in vascular-targeted interventions.

2.6. In vivo therapeutic efficacy

To validate localized NO release at the lesion site, we quantified NO levels in aortic plaques and non-plaque vascular regions using the Griess assay. As shown in Fig. S10A, a significant increase in NO concentration was observed in the atherosclerotic plaques of the B-NPs@MM + US group, while minimal changes were found in non-lesion regions (Fig. S10B), supporting site-specific NO generation. These findings confirm the targeted NO delivery capability of the platform, consistent with its therapeutic efficacy in vivo.

Building upon the encouraging in vitro outcomes, we further validated the therapeutic efficacy of B-NPs@MM in ApoE^−/− mice fed a high-fat diet (HFD) to mimic early-stage AS. As illustrated in Fig. 6A, mice underwent three consecutive US-triggered treatments starting from week 2 of HFD, and evaluations were performed at week 12. Gross aortic observation (Fig. 6B and Fig. S11) and Oil Red O staining (Fig. 6C) revealed extensive plaque deposition in the AS group. Quantitative analysis (Fig. 6D) showed that B-NPs@MM + US treatment significantly reduced plaque burden compared with AS (P < 0.001), B-NPs@MM (P < 0.05), and B-NPs + US (P < 0.05) groups, suggesting a superior anti-atherosclerotic effect achieved through targeted, US-triggered NO release. We next assessed the systemic inflammatory status via serum IL-1β and TNF-α levels (Fig. 6E and F). B-NPs@MM + US markedly suppressed both cytokines relative to all other groups (P < 0.001), indicating effective inhibition of vascular inflammation.

Fig. 6.

In vivo evaluation of therapeutic efficacy of B-NPs@MM in ApoE^−/− mice. (A) Schematic illustration of the animal experiment. ApoE^−/− mice were fed a high-fat diet (HFD) for a total of 12 weeks. (B) Gross images of the aortic arch and root area after treatment, with atherosclerotic plaques indicated by black arrows. (C) Representative en face Oil Red O staining of entire aortas showing lipid-rich plaques (red). (D) Quantification of Oil Red O-positive lesion area as a percentage of total aortic surface area. (E–F) ELISA analysis of serum pro-inflammatory cytokines IL-1β (E) and TNF-α (F) after treatment.

Histological analysis further supported these findings. Cell proliferation, a hallmark of lesion progression, was assessed by Ki67 staining (Fig. 7A and B). The AS group exhibited the highest proportion of Ki67⁺ cells, indicating robust proliferative activity within atherosclerotic lesions. While B-NPs@MM and B-NPs + US moderately reduced Ki67⁺ cell proportions (P < 0.01 vs AS), the B-NPs@MM + US group demonstrated the most significant reduction (P < 0.0001 vs AS), highlighting the inhibitory effect of US-triggered delivery on pathological cell proliferation. The difference between B-NPs + US and B-NPs@MM + US also reached statistical significance (P < 0.01), suggesting the additional benefit of macrophage membrane-mediated targeting. Macrophage infiltration, as marked by CD68 immunostaining (Fig. 7C and D), was abundant in the AS group and reduced in all treatment groups. Notably, B-NPs@MM + US led to the lowest proportion of CD68⁺ cells (P < 0.001 vs AS; P < 0.01 vs B-NPs@MM or B-NPs + US), indicating a pronounced anti-inflammatory effect potentially attributed to enhanced NO bioavailability and precise lesion targeting. Matrix metalloproteinase-9 (MMP-9) is a key enzyme involved in extracellular matrix degradation and plaque destabilization [46]. As shown in Fig. 7E and F, MMP-9⁺ cell abundance was significantly lower in the B-NPs@MM + US group compared to all other groups (P < 0.001 vs AS, P < 0.05 vs B-NPs + US), suggesting that treatment suppressed ECM degradation and may contribute to plaque stabilization. In contrast, CD31, an endothelial marker reflecting re-endothelialization and vascular repair [47], was significantly elevated in the B-NPs@MM + US group (Fig. 7G and H). Compared to AS, B-NPs@MM, and B-NPs + US groups, the B-NPs@MM + US treatment restored endothelial coverage more effectively (P < 0.001 vs AS; P < 0.05 vs B-NPs + US), aligning with prior in vitro findings on endothelial regeneration.

Fig. 7.

Immunohistochemical analysis of plaque cellular activities and microenvironmental changes in the aortic root. (A) Representative images of Ki67 immunostaining with magnified views. (B) Quantification of Ki67⁺ proliferating cells as a percentage of total cells. (C) Representative images of CD68 immunostaining with magnified views. (D) Quantification of CD68⁺ macrophages. (E) Representative images of MMP-9 immunostaining with magnified views. (F) Quantification of MMP-9⁺ cells. (G) Representative images of CD31 immunostaining with magnified views. (H) Quantification of CD31⁺ endothelial cells.

In Fig. 8, we further evaluated smooth muscle cells (SMCs) content using α-SMA staining (Fig. 8A and B). α-SMA⁺ SMCs are a critical component of fibrous caps that contribute to plaque stability [48]. The B-NPs@MM + US group showed a significant increase in α-SMA⁺ cell density (P < 0.0001 vs AS; P < 0.01 vs B-NPs + US), indicating better maintenance of fibrous cap integrity. Masson's trichrome staining was employed to assess collagen content, another hallmark of stable plaques (Fig. 8C and D). Collagen deposition was markedly enhanced in the B-NPs@MM + US group compared with all other groups (P < 0.0001 vs AS; P < 0.05 vs B-NPs + US), reinforcing the therapeutic benefits in promoting fibrous plaque remodeling. Finally, HE staining was performed to visualize overall plaque structure and morphology, including lipid core size, intimal thickening, and cell infiltration (Fig. 8E). Compared to AS and control treatments, the B-NPs@MM + US group exhibited smoother luminal surfaces, thinner neointima, and reduced necrotic core areas, collectively indicative of plaque regression and structural stabilization.

Fig. 8.

Histological and immunohistochemical evaluation of plaque remodeling and stability.(A) Representative immunohistochemical images of α-SMA staining in the aortic root, with magnified views showing smooth muscle cell distribution.(B) Quantification of α-SMA⁺ cells expressed as a percentage of total cells.(C) Representative images of Masson trichrome staining for collagen fibers (blue), with corresponding magnified regions.(D) Quantification of collagen content as a percentage of total plaque area.(E) Representative hematoxylin and eosin (H&E) staining images of aortic root sections, with magnified views. Scale bars = 300 μm.

Collectively, these in vivo results demonstrate that B-NPs@MM combined with US stimulation effectively suppresses plaque progression, mitigates inflammation, promotes endothelial repair, and alleviates fibrotic remodeling in atherosclerotic lesions. These therapeutic benefits are not solely attributed to NO's direct vasoprotective effect, but also stem from its multifaceted roles in reducing inflammation, restoring endothelial homeostasis, and inhibiting VSMC proliferation—key elements in the early pathogenesis of atherosclerosis. Importantly, this therapeutic strategy exerts maximal efficacy during the early stages of AS, particularly when endothelial dysfunction first emerges. Our findings are consistent with the growing body of evidence supporting the use of nanotechnology-based therapies to modulate vascular pathology and inflammation in cardiovascular diseases [6]. Therefore, US-triggered, targeted NO delivery offers a promising avenue for early clinical intervention in vascular disease management.

2.7. Biosafety assessment

To evaluate the in vivo biocompatibility of B-NPs@MM, C57BL/6 mice were intravenously injected with B-NPs, B-NPs@MM, or their US-irradiated counterparts. Histological examination of major organs by HE staining showed no signs of tissue damage, inflammation, or necrosis in the heart, liver, spleen, lungs, or kidneys across all groups (Fig. 9A). The integrity of cardiac fibers, hepatic lobules, pulmonary alveoli, splenic architecture, and renal tubules was well preserved. Additionally, serum were collected for biochemical and histological analysis. As shown in Fig. 9B–E, serum biochemical parameters including aspartate aminotransferase (AST), alanine aminotransferase (ALT), creatinine (CREA-S), and urea (UREA) remained within normal physiological ranges in all treatment groups. No significant elevation was observed, indicating no evident hepatic or renal toxicity induced by the nanoparticles or US stimulation. These findings suggest that both B-NPs and B-NPs@MM exhibit favorable systemic safety profiles under experimental conditions. Collectively, the results indicate that B-NPs@MM, with or without US exposure, does not elicit significant organ toxicity and is biocompatible for potential in vivo therapeutic applications.

Fig. 9.

In vivo biosafety evaluation of B-NPs@MM in ApoE^−/− mice. (A) Representative H&E-stained histological sections of major organs (heart, liver, spleen, lung, kidney) from normal control (NC), AS, and treatment group. (B–E) Blood biochemical analysis of liver and kidney function indicators, including (B) aspartate aminotransferase (AST), (C) alanine aminotransferase (ALT), (D) creatinine (CREA-S), and (E) urea nitrogen (UREA), after various treatments.

Ex vivo fluorescence imaging (Fig. S12) revealed that B-NPs@MM showed significantly reduced hepatic accumulation compared to uncoated B-NPs, while distribution in other major organs remained comparable. This reduction is likely due to the CD47-mediated immune evasion conferred by the macrophage membrane, which helps B-NPs@MM avoid rapid clearance by the mononuclear phagocyte system and improves their in vivo delivery profile [38].

As shown in Fig. S13, no statistically significant differences were observed among AS, B-NPs@MM, B-NPs + US, and B-NPs@MM + US groups in total cholesterol (TC), low-density lipoprotein cholesterol (LDL-C), high-density lipoprotein cholesterol (HDL-C), or triglycerides (TG) levels. These findings indicate that the therapeutic intervention did not notably affect systemic lipid metabolism during the treatment window. The observed atheroprotective effects were therefore primarily attributable to local anti-inflammatory, endothelial protective, and anti-proliferative actions rather than lipid-lowering effects.

To evaluate systemic safety, systolic blood pressure (SBP) was monitored in both healthy and atherosclerotic mice following nanoparticle administration. As shown in Fig. S14, no significant SBP fluctuations were observed in any group, including B-NPs@MM + US, over the 3-day period. Despite NO's vasodilatory properties, the localized delivery and moderate release dose did not induce systemic hypotension. These results, consistent with our biosafety findings, confirm that B-NPs@MM-mediated NO delivery operates within a safe therapeutic window.

3. Conclusion

In this study, we developed a biomimetic nanomedicine platform—a macrophage membrane-camouflaged nanoparticle (B-NPs@MM)—capable of US-triggered NO release for targeted early intervention of AS. This nanosystem features a core-shell architecture, where the NO donor BNN6 is encapsulated within PLGA nanoparticles and cloaked with a macrophage-derived membrane, achieving immune evasion, lesion targeting, and on-demand NO release under external US stimulation. Comprehensive in vitro and in vivo evaluations demonstrated that B-NPs@MM + US significantly promotes endothelial cell proliferation, migration, and angiogenesis, while concurrently inhibiting VSMC proliferation and reducing oxidative stress and inflammation. These functions are crucial for maintaining endothelial integrity, restoring vascular homeostasis, and preventing early atherogenesis.

Collectively, this work highlights a novel nano-macrophage-based strategy for precise, controllable NO delivery, which effectively modulates key cellular events in early-stage AS. It offers a promising therapeutic paradigm for noninvasive, targeted vascular repair and clinical translation.

4. Materials and methods

4.1. Synthesis and characterization of BNN6

The synthesis of BNN6 was performed according to our previously reported procedure [49]. Specifically, 2.34 mL (10 mmol) of N,N′-bis-sec-butylamino-p-phenylenediamine (BPA, TCI America) was dissolved in 18 mL of absolute ethanol under continuous stirring. Under a nitrogen atmosphere, 20 mL of degassed 6 M sodium nitrite (NaNO₂, Sigma-Aldrich) aqueous solution was added to the reaction mixture. After stirring for 30 min, 20 mL of 6 M hydrochloric acid solution was introduced dropwise using a separating funnel. During the reaction, the solution color gradually shifted from orange to red, and a beige precipitate began to form. After 4 h of reaction at room temperature, the suspension was centrifuged at 1000 rpm for 15 min to isolate the solid product. The precipitate was washed three times with deionized water and 50 % ethanol–water (v/v) mixture to remove unreacted reagents, and then dried overnight under vacuum in the dark. The resulting BNN6 product was characterized by high-resolution mass spectrometry (Orbitrap LC/MS, Thermo Fisher Scientific).

4.2. Isolation of macrophage membrane

Macrophage membranes were isolated from RAW264.7 cells according to a previously reported protocol with minor modifications [27]. Briefly, RAW264.7 cells were harvested and suspended in a membrane protein extraction buffer provided by a commercial membrane protein isolation kit (Beyotime, China). The cell suspension was incubated on ice for 15 min to initiate membrane disruption. Subsequently, the cells were transferred to a glass homogenizer and mechanically homogenized with approximately 30 strokes. The homogenate was first centrifuged at 1500 rpm for 10 min at 4 °C to remove unbroken cells and nuclei. The supernatant was then collected and centrifuged at 14,000 rpm for 30 min at 4 °C to pellet the membrane fraction. The collected membrane pellet was resuspended in cold PBS and the total protein concentration was determined using a bicinchoninic acid (BCA) protein assay kit (Thermo Fisher Scientific, USA). To prepare membrane-derived vesicles, the obtained macrophage membranes were subjected to ultrasonication for 15 min in an ice bath, followed by extrusion through a 400 nm polycarbonate porous membrane using a mini-extruder (Avestin LF-1, Canada) for 10 cycles. The resulting membrane vesicles (MM) were stored in deionized water at 4 °C for further use.

4.3. Preparation and characterization of B-NPs and B-NPs@MM

BNN6-loaded PLGA nanoparticles (B-NPs) were prepared using a standard oil-in-water (O/W) emulsion solvent evaporation method with slight modifications [21]. Briefly, 2 mg of BNN6 and 10 mg of PLGA were dissolved in 500 μL of dichloromethane. The organic phase was added dropwise into 3 mL of 0.5 % (w/v) polyvinyl alcohol (PVA) aqueous solution and emulsified using a probe sonicator (LC1000N, Ultrasonic Processor, China) for 5 min under ice-bath conditions to form a stable O/W emulsion. The resulting emulsion was then transferred into 10 mL of deionized water and stirred gently in a fume hood for 3 h to allow complete solvent evaporation. The excess BNN6 was collected and its absorption at 380 nm was measured to calculate the entrapment and loading efficiency of the carriers. The nanoparticles were collected by centrifugation at 20,000×g for 20 min and washed twice with deionized water. For fluorescent labeling, DiI or DiR was added to the organic phase at 0.1 wt% relative to PLGA prior to emulsification.

To fabricate macrophage membrane-coated B-NPs (B-NPs@MM), an appropriate amount of previously isolated macrophage membrane vesicles was added to the freshly prepared B-NPs at a membrane-to-particle protein-to-polymer weight ratio of 1:1 [50,51]. The mixture was subjected to sonication in a bath sonicator (KH-160TDB, KUNSHAN, China) for 5 min to promote membrane fusion and coating. The resulting B-NPs@MM were collected by centrifugation at 20,000×g for 20 min and resuspended in PBS for further use.

The hydrodynamic diameter, polydispersity index (PDI), and surface zeta potential of B-NPs and B-NPs@MM were measured using a DLS instrument (NanoBrook 90 Plus PALS, Brookhaven Instruments, USA). The successful membrane coating was confirmed by TEM (TEM, Hitachi HT7700, Japan) operated at 100 kV. FT-IR spectroscopy (Nicolet iS50R, Thermo Scientific, USA) was performed to analyze characteristic chemical bonds and membrane integration over a spectral range of 400–4000 cm⁻¹. TGA was performed from room temperature to 500 °C at a heating rate of 10 °C/min under nitrogen atmosphere (TA Instruments, USA). Raman spectroscopy (LabRAM HR800) was performed to detect characteristic molecular vibrations over a spectral range of 100–2000 cm⁻¹.

4.4. Examination of macrophage membrane proteins

To verify the successful coating of macrophage membranes onto B-NPs, protein profiles were analyzed using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Western blotting. Four sample groups were prepared: whole macrophage cells (RAW264.7), isolated macrophage membranes (MM), uncoated B-NPs, and membrane-coated nanoparticles (B-NPs@MM). Each sample (50 μL, 15 μg/mL in total protein content) was centrifuged and resuspended in 20 μL of loading buffer under reducing conditions to solubilize membrane-associated proteins. A 5 μL aliquot of each sample was loaded onto a 10 % SDS-PAGE gel and run at 100 V for 100 min using a vertical electrophoresis system (VE 680, Tanon, China). After electrophoresis, protein bands were visualized using silver staining.

For Western blot analysis, proteins were transferred onto polyvinylidene fluoride (PVDF) membranes following SDS-PAGE and blocked with 5 % non-fat milk in TBST for 1 h at room temperature. Membranes were then incubated overnight at 4 °C with primary antibodies against macrophage membrane-specific markers, including CD11b, integrin α4, and CD47. After washing, membranes were incubated with appropriate HRP-conjugated secondary antibodies for 1 h at room temperature. Protein bands were detected using enhanced chemiluminescence (ECL) reagents and imaged with a gel imaging system.

4.5. In vitro US parameters exploration and NO detection

NO release was quantified using the Griess assay Kit (Beyotime, China), which detects the stable oxidation products of NO, namely nitrite (NO₂⁻) and nitrate (NO₃⁻). Therefore, the values represent total NO metabolites rather than free NO. Although it does not reflect real-time NO flux, the cumulative levels of NO₂⁻/NO₃⁻ are commonly used as surrogate indicators for NO bioavailability in vitro and in vivo [52,53].

BNN6 (100 μM), B-NPs, and B-NPs@MM dispersions (equivalent to 100 μM BNN6) were prepared in PBS and subjected to US irradiation using a gene transfection US device (Nepa Gene, Japan) under the following parameters: 1.0 W/cm² intensity, 50 % duty cycle, and 1 MHz frequency. Each sample (1 mL) was exposed to US for cumulative time periods of 0, 10, 20, 30, 40, and 60 min at room temperature. At each designated time point, 50 μL of the sample was withdrawn and added to a 96-well plate, followed by the addition of 50 μL of Griess Reagent I and 50 μL of Griess Reagent II. After 10 min of incubation in the dark at room temperature, the absorbance was measured at 540 nm using a microplate reader (TECAN, Switzerland). Sodium nitrite (NaNO₂) was used to generate the standard calibration curve. Samples without US treatment served as controls.

4.6. Haemocompatibility assay

The hemocompatibility of B-NPs and B-NPs@MM was assessed using a standard hemolysis assay. Fresh mouse blood was collected with EDTA and centrifuged at 1000×g for 5 min to isolate erythrocytes. The erythrocytes were washed three times with sterile PBS and resuspended to a final hematocrit of 2 %. Then, 200 μL of the erythrocyte suspension was incubated with 800 μL of B-NPs or B-NPs@MM dispersion at various concentrations (typically 100–500 μg/mL) for 2 h at 37 °C. Saline and 1 % Triton X-100 served as negative and positive controls, respectively. After incubation, samples were centrifuged at 1000×g for 5 min, and the absorbance of the supernatant was measured at 541 nm using a microplate reader. Hemolysis percentage was calculated using the formula: Hemolysis (%) = Apositive−Anegative Asample−Anegative × 100 %

4.7. Cell viability

The cytocompatibility of B-NPs and B-NPs@MM was evaluated using a Cell Counting Kit-8 (CCK-8, Dojindo, Japan) following the manufacturer's instructions. HUVECs, VSMCs, and RAW264.7 cells were seeded in 96-well plates (5 × 10³ cells/well) and treated with B-NPs or B-NPs@MM at BNN6-equivalent concentrations of 0.05–800 μM for 24 or 48 h in serum-free DMEM. After treatment, cells were washed with PBS and incubated with CCK-8 solution (10 % in fresh medium) for 2 h at 37 °C. Absorbance at 450 nm was recorded using a microplate reader (TECAN) to assess cell viability.

4.8. Evaluation of targeting efficiency in vitro

To evaluate cellular targeting capability, HUVECs and VSMCs were seeded in confocal dishes and pretreated with hydrogen peroxide (H₂O₂, 100 μM) for 4 h to simulate an inflammatory microenvironment. After washing with PBS, cells were incubated with DiI-labeled B-NPs or B-NPs@MM (equivalent to 100 μM BNN6) for 1 h at 37 °C under normoxic conditions. Following incubation, cells were rinsed thoroughly with PBS to remove unbound nanoparticles. Nuclei were counterstained with DAPI for 5 min, and cells were imaged using a fluorescence microscope (Olympus, Japan). Untreated cells were used as controls to assess basal uptake levels.

4.9. EdU proliferation assay

To simulate endothelial injury, HUVECs or VSMCs were pretreated with hydrogen peroxide (H₂O₂, 100 μM) for 2 h. After washing, cells were incubated with either B-NPs or B-NPs@MM (equivalent to 100 μM BNN6) that had been pre-irradiated with US for 10 min (1.0 W/cm², 50 % duty cycle, 1 MHz). Parallel groups without US irradiation served as controls. All cells were cultured for an additional 4 h in a CO₂ incubator at 37 °C before further analysis. Cell proliferation was evaluated using EdU Proliferation Kit (Beyotime, China) according to the manufacturer's protocol. Briefly, after nanoparticle treatment, HUVECs were incubated with 10 μM EdU for 2 h, fixed with 4 % paraformaldehyde, permeabilized with 0.5 % Triton X-100, and stained with Apollo fluorescent dye. Nuclei were counterstained with DAPI. EdU-positive cells were imaged by fluorescence microscopy and analyzed using ImageJ.

4.10. Transwell migration assay

Cells were treated as described above. Transwell chambers with an 8 μm pore size (Corning) were used to assess cell migration. After treatment, HUVECs were seeded in the upper chamber (2 × 10⁴ cells/well) in serum-free medium. The lower chamber was filled with DMEM containing 10 % FBS as a chemoattractant. After 12 h, non-migrated cells on the upper side were removed. Migrated cells on the lower surface were fixed with 4 % paraformaldehyde and stained with 0.1 % crystal violet. After imaging, crystal violet was extracted using 33 % acetic acid, and absorbance was measured at 595 nm to quantify cell migration.

4.11. Tube formation assay

Cells were treated as described above. Pre-cooled 48-well plates were coated with 100 μL/well of growth factor-reduced Matrigel (Corning, USA) and incubated at 37 °C for 30 min. HUVECs (passage 3–5) were then seeded at 2 × 10⁴ cells/well and incubated with treated or control nanoparticles (B-NPs, B-NPs@MM, ±US) during the reoxygenation period. After 6 h, tube-like structures were imaged using an inverted microscope (Olympus, Japan). Quantitative analysis of total tube length and branching points was performed using the Angiogenesis Analyzer plugin in ImageJ.

4.12. Scratch wound healing assay

Cells were treated as described above. HUVECs were grown to confluence in 6-well plates and serum-starved for 12 h prior to scratching. A straight wound was created using a sterile 200 μL pipette tip, and detached cells were removed by PBS washing. Cells were then treated with B-NPs or B-NPs@MM (±US) in serum-free medium. Wound closure was imaged at 0 h and 24 h using a microscope. The percentage of wound closure was quantified using ImageJ software.

4.13. DAF-FM DA fluorescence detection

HUVECs were pretreated with H₂O₂, and RAW264.7 cells were stimulated with LPS to simulate oxidative and inflammatory conditions, respectively. Cells were then incubated with or without B-NPs@MM and exposed to US. Where indicated, L-NAME (100 μM) or 1400W (100 μM) was added 1 h prior to stimulation to inhibit eNOS or iNOS activity. After US activation, cells were stained with DAF-FM DA (5 μM) for 30 min in the dark, and fluorescence intensity was analyzed under a fluorescence microscope and quantified using ImageJ.

4.14. Measurement of cGMP levels

To evaluate the bioactivity of released NO, intracellular cGMP levels were quantified using a commercially available cGMP ELISA kit according to the manufacturer's instructions. Briefly, HUVECs were lysed on ice using 0.1 M pre-chilled hydrochloric acid (HCl), followed by incubation for 15 min to ensure complete lysis. Lysates were centrifuged at 12,000 g for 10 min at 4 °C, and supernatants were transferred into pre-chilled tubes. A 50 μL aliquot of each sample was subjected to ELISA analysis. Absorbance was measured at 450 nm using a microplate reader, and cGMP concentrations were calculated based on a standard curve. (n = 3).

4.15. Flow cytometry analysis of nanoparticle-cell interaction

To evaluate the cellular interaction of nanoparticles with macrophages and endothelial cells, DiD-labeled B-NPs and B-NPs@MM were incubated with RAW264.7 and HUVEC cells, respectively. Briefly, cells were seeded in 6-well plates at a density of 2 × 10⁵ cells/well and cultured overnight. DiD-labeled nanoparticles were then added to the medium at an equivalent DiD concentration and incubated for 1 h at 37 °C. After incubation, cells were washed three times with PBS to remove unbound particles, detached using trypsin-EDTA (for HUVECs) or cell scraper (for RAW264.7), and resuspended in cold PBS. The percentage of DiD-positive cells was quantified using a BD Accuri™ C6 flow cytometer, and data were analyzed with FlowJo software. Blank cells without nanoparticle treatment served as negative controls.

4.16. Animal models and ethical considerations

Male C57BL/6 mice and apolipoprotein E knockout (ApoE−/−) mice (6-week-old) were procured from Cyagen Biosciences under specific pathogen-free conditions. After two-week acclimatization with standard chow, ApoE−/− mice (8-week-old) received atherogenic induction through 12-week feeding with customized high-fat diet (20 % fat, 1.25 % cholesterol). All experimental protocols were approved by the Ethics Committee of Southwest Medical University (Approval No.: 20230812-008).

4.17. Targeted delivery evaluation

To assess the in vivo targeting capability of the B-NPs@MM nanoparticle, atherosclerotic ApoE−/− mice (n = 3) were intravenously injected via the tail vein with 100 μL of DiR-labeled nanoparticles (2 mg/mL), including both uncoated B-NPs and macrophage membrane-camouflaged B-NPs@MM as experimental and control groups, respectively. After 24 h of systemic circulation, the mice were anesthetized with intraperitoneal injection of sodium pentobarbital (50 mg/kg) and subsequently perfused through the left ventricle with 20 mL of heparinized saline to remove residual blood and reduce background fluorescence. Following perfusion, major organs including the heart, liver, spleen, lungs, kidneys, as well as the entire aortic tree were carefully excised and rinsed in cold PBS. Ex vivo fluorescence imaging was performed using the IVIS Spectrum imaging system (PerkinElmer, USA) under standardized exposure conditions (excitation: 745 nm; emission: 800 nm). Fluorescence intensity was quantitatively analyzed using Living Image software to compare nanoparticle accumulation across different tissues.

4.18. Measurement of NO levels in plaque and non-plaque vascular regions

To quantify local NO production, aortic tissues from ApoE^−/− mice were dissected after treatment, and both plaque-rich (aortic arch and abdominal regions) and non-plaque vascular segments were isolated. Tissues were immediately homogenized in PBS with protease inhibitors, and centrifuged at 12,000 rpm for 10 min at 4 °C. The supernatant was collected, and nitrite concentration—a stable NO metabolite—was measured using a Griess Reagent Kit (Beyotime, China), following the manufacturer's instructions. Total protein concentration was quantified by BCA assay, and NO levels were normalized to protein content (nmol/mg protein). All samples were assayed in triplicate (n = 4).

4.19. Therapeutic intervention protocol

After 2 weeks of dietary induction, ApoE−/− mice were randomly allocated into five groups (n = 6): 1) PBS control; 2) B-NPs@MM (10 mg/kg, weekly IV); 3) B-NPs + US (nanoparticles with US); 4) B-NPs@MM + US. US irradiation (1 MHz, 1 W/cm²) was applied for 3 consecutive days post-injection. Treatment continued for 10 weeks prior to terminal analysis.

4.20. Systemic biomarker profiling

Blood specimens underwent comprehensive analysis using standardized clinical protocols. Lipid profiles including TC, TG, HDL-C, and LDL-C were quantified through enzymatic colorimetric assays (Roche Diagnostics). Concurrently, proinflammatory cytokines interleukin-1β (IL-1β) and tumor necrosis factor-α (TNF-α) concentrations were determined via commercial ELISA kits (R&D Systems) following manufacturer specifications, with optical density measurements conducted at 450 nm using a microplate reader (BioTek Synergy H1).

4.21. Histopathological assessment

At the predetermined endpoint, mice were deeply anesthetized with pentobarbital (50 mg/kg) and euthanized via cervical dislocation. The entire aortic tree, from the aortic arch to the iliac bifurcation, was carefully dissected, rinsed with cold PBS, and fixed in 4 % paraformaldehyde overnight at 4 °C. For gross plaque assessment, en face Oil Red O staining was performed on longitudinally opened aortas to visualize lipid-rich lesions. The percentage of lesion area was quantified using ImageJ software and expressed as a proportion of the total aortic surface area. For cross-sectional plaque analysis, hearts containing the aortic root were embedded in optimal cutting temperature (OCT) compound and cryosectioned. Serial 10-μm-thick frozen sections were obtained at 100-μm intervals starting from the appearance of the aortic valve leaflets. Selected sections were subjected to hematoxylin and eosin (H&E) staining for general morphology and Masson's trichrome staining to assess collagen content and fibrous cap structure. Immunohistochemical analyses were performed on adjacent sections using primary antibodies specific to key cellular and molecular markers of plaque biology: CD68 (macrophages), α-smooth muscle actin (α-SMA; vascular smooth muscle cells), CD31 (endothelial cells), matrix metalloproteinase-9 (MMP-9; extracellular matrix degradation), and Ki67 (proliferation marker). Visualization was achieved using DAB-based chromogenic detection, and positively stained areas or cells were quantified using standardized thresholds and ROI settings in ImageJ.

4.22. Biosafety profiling

To evaluate the systemic biosafety of the B-NPs@MM platform, major hematological and histological parameters were examined post-treatment. At the study endpoint, whole blood was collected via cardiac puncture into EDTA-coated tubes and immediately subjected to complete blood count (CBC) analysis using an automated hematology analyzer (Sysmex XN-1000, Japan) to assess potential hematological toxicity. Serum was separated by centrifugation at 3000 rpm for 10 min and analyzed for hepatic and renal function markers, including ALT, AST, CREA-S, and UREA, using commercially available assay kits (Roche Diagnostics, Germany) following the manufacturer's protocols. For histopathological evaluation, major organs including the heart, liver, spleen, lungs, and kidneys were harvested, fixed in 4 % paraformaldehyde, embedded in paraffin, sectioned at 5 μm, and stained with hematoxylin and eosin (H&E). Tissue sections were examined under a light microscope to identify signs of inflammation, necrosis, or other pathological changes indicative of organ toxicity. In addition, SBP was measured on days 0,1, 2, and 3 post-injection using a non-invasive tail-cuff system (BP-2010A, Softron, Beijing, China; n = 4) to monitor potential systemic vascular effects following NO release.

4.23. Statistical analysis

Graphs were plotted and appropriate statistical analyses were conducted using GraphPad Prism 9 and OriginPro 9. All quantitative results are expressed as the mean ± SD. Statistical analysis was performed by Student's t-test (α = 0.05) or one-way analysis of variance (ANOVA). Statistical significance was set at ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001.

CRediT authorship contribution statement

Lingling Xu: Writing – original draft, Methodology, Investigation, Formal analysis, Data curation. Daifang Zhang: Methodology. Li Song: Methodology. Yifei Wu: Methodology. Longqi Jiang: Methodology. Zhenyu Liu: Methodology. Xin Qian: Methodology. Jun Zhou: Writing – review & editing, Supervision. Yong Liu: Writing – review & editing, Supervision. Ya Wu: Visualization, Validation, Supervision, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

The following is/are the supplementary data to this article.

Data availability

Data will be made available on request.