Ultrasound-Responsive Dual-Prodrug Nanoassembly for "Fenestrae-Restoration Strategy" in Liver Fibrosis Therapy.

1
Introduction
Chronic liver diseases and prolonged, recurrent liver injury can lead to liver fibrosis, a condition characterized by excessive deposition of extracellular matrix (ECM), parenchymal cell death, and persistent chronic inflammation [1, 2, 3, 4]. Without effective intervention at the liver fibrosis stage, the disease may progress to cirrhosis, which is a leading cause of death worldwide [5]. Since activated hepatic stellate cells (aHSCs) are the primary source of ECM deposition, numerous anti‐fibrotic strategies have focused, through diverse mechanistic approaches, on suppressing HSCs activation [6, 7, 8]. However, in the fibrotic environment, liver sinusoidal endothelial cells (LSECs) act as a barrier between the bloodstream and HSCs, impeding the transport of therapeutic agents, including nanomedicines. This barrier represents a largely overlooked obstacle to efficient anti‐fibrotic therapy [9, 10]. Therefore, any delivery system targeting HSCs in the space of Disse must first overcome the LSECs barrier.
LSECs are the most abundant non‐parenchymal cells in the liver. LSECs are a specialized class of endothelial cells that form a permeable barrier between the liver sinusoids and both HSCs and hepatocytes. Under physiological conditions, LSECs form unique sinusoidal capillary channels and exhibit a distinctive fenestrated phenotype, characterized by pores known as “fenestrae”. These fenestrae, with ∼50–200 nm in diameter, cluster together to form “sieve plates”, creating open portals to facilitate bidirectional exchange of solutes, oxygen, and metabolites between the sinusoidal lumen and the space of Disse [11]. In a healthy liver, differentiated LSECs possess abundant fenestrae and a lack of basement membrane, which enhances the efficient exchange of materials and facilitates the direct delivery of therapeutic agents to HSCs [12, 13]. In contrast, during liver fibrosis, LSECs undergo capillarization, a process marked by the progressive loss or complete disappearance of fenestrae associated by development of a continuous basement membrane. This transformation reduces the porosity of LSECs, significantly impeding the exchange of substances between the bloodstream and the space of Disse, thereby limiting drug uptake and decreasing accumulation of therapeutic agents in HSCs [14, 15]. In vitro studies have shown that healthy, differentiated LSECs inhibit HSCs activation and promote the reversion of aHSCs to quiescent HSCs (qHSCs), whereas capillarized LSECs lose this regulatory capacity. In cirrhotic experimental models, therapeutic strategies aimed at improving vascular tone by targeting LSECs have been shown to alleviate liver fibrosis effectively [16]. Notably, LSECs capillarization is an early event that precedes both HSCs activation and the onset of liver fibrosis, thereby offering valuable insights into disease progression and opportunities for novel early therapeutic interventions [13, 16, 17].
Restoration of fenestrae, therefore, represents a highly promising therapeutic strategy, with nitric oxide (NO) playing a pivotal role in maintaining fenestrae integrity in LSECs. In fibrotic liver, capillarized LSECs exhibit diminished NO levels and reduced bioavailability, thereby losing their ability to sustain HSCs quiescence [14, 18, 19]. As a gaseous molecule, NO is a key vasodilator involved in regulating hepatic vascular tone; it readily crosses cell membranes and binds to its primary receptor, soluble guanylyl cyclase (sGC). Upon diffusion into cells, NO binds to the Fe2
+ in the heme group of sGC, activating its enzymatic activity and catalyzing the conversion of guanosine triphosphate (GTP) into the second messenger cyclic guanosine monophosphate (cGMP). This, in turn, activates downstream signaling pathways, including protein kinase G (PKG), ultimately leading to fenestrae restoration [18]. In recent years, the anti‐fibrotic effects of sGC stimulators have been demonstrated not only in the cardiovascular system but also in other organs such as the kidney, skin, and lung [9, 20, 21, 22]. A variety of NO donors have been developed, including S‐nitrosothiols, N‐nitrosamines, metal NO complexes, bis‐N‐nitroso compounds (BNNs), and others [23]. Among these, S‐nitrosothiols (RSNOs), S‐nitroso derivatives of thiols, have been reported to exhibit ultrasound (US)‐responsive NO release. This is advantageous, since, as a preferred exogenous stimulus, US is non‐invasive and offers deep tissue penetration, enabling precise energy delivery to deep‐seated targets such as the liver. Importantly, US can be turned on and off with high temporal precision, rendering on‐demand NO release. Moreover, by adjusting the transducer position and acoustic parameters, the delivered energy can be highly confined to the hepatic region, thereby minimizing unintended effects on surrounding tissues.
Metformin (Met), a classic example of the “old drug, new tricks” paradigm, has recently emerged as a potent agent against both tumor growth and fibrogenesis [24, 25]. Mechanistically, Met activates AMP‐activated protein kinase (AMPK), thereby inhibiting mammalian target of rapamycin (mTOR) and its downstream effector phosphoprotein 70 ribosomal protein S6 kinase (p70S6K), leading to the suppression of HSCs activation [26, 27, 28]. However, due to the asynchronous pharmacokinetic (PK) profiles of NO prodrugs and free Met, systemic administration of unformulated Met is burdened by unpredictable PK behavior, dose‐limiting toxicity, and low bioavailability [29]. Inspired by the seminal work of Leaf Huang [25], we therefore engineered a micelle‐forming Met prodrug (PMet). Owing to its amphiphilic architecture, PMet self‐assembles into stable micelles that not only enhance structural integrity but also leverage the intrinsic guanidinium motif to improve cellular uptake [30, 31].
Given the critical roles of LSECs and HSCs in the progression of liver fibrosis, we developed an US‐responsive dual‐prodrug polymeric nanoassembly (PMS), composed of a NO prodrug (mSNO) and PMet (Figure 1A). Upon US exposure, US‐induced cavitation promotes the low‐energy S─N bond cleavage in the ─SNO moiety, leading to a controlled release of NO [32, 33, 34, 35]. Acting as a potent sGC stimulator, NO reprograms capillarized LSECs back to a differentiated phenotype via activation of the NO‐sGC‐cGMP signaling pathway. This process restores fenestrae and disrupts the pathological barrier function of capillarized LSECs. The restoration of fenestrae then facilitates the enhanced delivery of PMet, allowing it to traverse the endothelial layer and reach the space of Disse. Once internalized, PMet undergoes hydrolysis by intracellular esterases, cleaving the C─C bond of PMet to release active Met. This activates AMPK, thereby downregulating the mTOR pathway and its downstream effector p70S6K. This leads to inhibition of the proliferation and migration of aHSCs, contributing to remodeling of the fibrotic liver microenvironment (Figure 1B). Overall, we propose this smart fenestrae‐restoring strategy not only enhances drug delivery and bioavailability but also represents a promising avenue for the development of advanced modalities for liver fibrosis.

2
Results and Discussion
2.1
Synthesis and Characterization of PMS
We first synthesized polyvinylbenzyl chloride (PVBC) via reversible addition‐fragmentation chain transfer (RAFT) polymerization (Figure S1a). The chemical structure of PVBC is shown in Figure 2a and was confirmed by 1H nuclear magnetic resonance (1H NMR) (Figure 2b). PVBC exhibits characteristic resonances consistent with the repeating units of 4‐vinylbenzyl chloride (VBC). Broad signals assigned to aromatic protons appear at δ 6.70–7.40 ppm (Ar–H), while the benzylic chloromethyl group (─CH2Cl) shows a signal at δ 4.40–4.70 ppm. In addition, the polymer backbone ─CH/─CH2 protons form a typical broad envelope in the δ 1.20–2.20 ppm region. Compared with the VBC monomer, the PVBC spectrum no longer displays the characteristic vinyl proton signals (─CH═CH2) in the δ 5.1–6.8 ppm range, indicating that the C═C double bond has been consumed and polymerization is complete. The presence of the key functional group resonances and the disappearance of the monomer vinyl signals provide strong evidence for the successful synthesis of PVBC, and the observed chemical shift ranges are in agreement with literature reports for PVBC [36]. Further confirmation of PVBC synthesis was obtained from Fourier‐transform infrared (FT‐IR) spectra (Figure 2c). The spectrum showed peaks associated with the benzene ring and methylene group at 3024, 1599, 1580, 1492, and 1449 cm−1. The peak at 835 cm−
1 indicated para‐substitution on the benzene ring. Additionally, peaks at 2920 cm−1 (O─H), 1733 cm−1 (C═O) confirmed the presence of carboxylic acid groups.
Next, PMet was prepared by chemically conjugating Met with PVBC. Transmission electron microscopy (TEM) revealed that PMet formed uniformly distributed spherical particles (Figure 2d). Compared to PVBC, PMet exhibited significantly improved water solubility (Figure S1b), attributed to the hydrophilic biguanide groups of Met. 1H NMR analysis showed the characteristic signal of the benzylic chloromethyl group (─CH2Cl) in PVBC at approximately δ 4.5 ppm (gray area) disappeared after the reaction, indicating substitution at the chloromethyl site. New signals absent in PVBC emerge in PMet around δ 2.8–3.1 ppm (green area), which can be assigned to protons associated with the N─CH3 groups of Met, thereby providing direct evidence for the covalent incorporation of Met into the polymer side chains. Meanwhile, the aromatic proton signals (δ 6.6–7.4 ppm) remain consistent in both spectra, confirming that the polymer backbone structure is preserved. A weak sharp singlet observed at δ 5.37 ppm is attributed to residual dichloromethane (DCM, CH2Cl2) from the purification process (Figure 2e). X‐ray photoelectron spectroscopy (XPS) results further show that the Cl content in PMet decreased significantly, while the N content increased markedly. Peak deconvolution was performed for the characteristic spin–orbit split doublet in the Cl 2p region (binding energy around ∼200 eV), the Cl 2p signal is strongly suppressed in PMet, indicating the substitution of ─CH2Cl groups after Met conjugation and supporting the successful synthesis of PMet (Figure S1c–e). The presence of the biguanide group was verified by colorimetric test and UV–vis spectrophotometry. After mixing PMet or Met with a chromogenic agent, which was prepared by mixing equal volumes of 10% (w/v) sodium nitroprusside with 10% (w/v) potassium hexacyanoferrate (III) and 10% sodium hydroxide, the solution color changed from yellow to red (Figure 2f). The intensity of the color correlated with the biguanide, whereas PVBC, due to its poor solubility, did not produce a visible color change. Partially undissolution of PMet was observed likely due to reduced solubility in the strongly alkaline conditions introduced by the chromogenic agent. UV–vis spectra (Figure S1f) revealed characteristic absorption peaks at 233 nm for both PMet and Met, associated with the biguanide group, which was not observed for PVBC. Additionally, PMet showed a low critical micelle concentration (CMC) value of 0.972 ± 0.543 µg mL−1 (Figure S1g), indicating good self‐assembly capability. The molecular weights of PVBC and PMet were ∼6.836 kDa and 6.206 kDa (Figure S1h), together with the rest of the data, suggesting well‐defined and controllable polymer structures. According to UV–vis spectrophotometry, the PMet loading capacity was 2.43 ± 0.05%.
In subsequent experiments, an NO prodrug mSNO was synthesized by reacting the thiol functional group of mPEG‐SH with TBN to produce mSNO (Figure 2g). FT‐IR spectra of mSNO exhibited characteristic peaks at 1525 and 775 cm−
1, corresponding to ─N═O and ─S ─N═O bonds, respectively (Figure 2h). UV–vis spectra also showed a distinct absorption peak at ∼325 nm, further confirming the presence of the ─SNO group (Figure 2i). 1H NMR spectra confirmed the disappearance of the ─SH peak at δ 1.86 ppm, indicating successful conversion of mPEG‐SH to mSNO (Figure 2j).
PMS was then formulated via electrostatic adsorption of mSNO onto PMet (Figure 2k). TEM image showed uniform spherical morphology (Figure 2l). Dynamic Light Scattering (DLS) and electrophoretic light scattering (ELS) revealed an average hydrodynamic diameter of 235 nm and a zeta potential of −12.9 mV (Figure S1i). As determined by UV–vis analysis, the electrostatic‐adsorption binding efficiency of PMet to mSNO reached 95.90 ± 0.90%. The UV–vis spectrum of PMS displayed absorption peak at 233 nm due to the biguanide group originating from PMet. However, the characteristic 325 nm peak of mSNO was less prominent, likely due to spectral overlap with PVBC (Figure S1f). FT‐IR spectra of PMS (Figure 2m) showed characteristic peaks of both PMet and mSNO, confirming successful electrostatic complexation. Importantly, no significant changes in hydrodynamic diameters were observed during 7 days of incubation in phosphate‐buffered saline (PBS, pH 7.4) containing 10% fetal bovine serum (FBS) at 37°C, indicating excellent colloidal stability (Figure S1j).
To evaluate the release properties of PMS, we examined both NO and Met release profiles. NO release was assessed under US exposure. Upon US irradiation, PMS released up to 20.6 ± 0.08 µm NO within 15 min (Figure 2n), as determined using a NaNO2 standard curve (Figure S1k). After US was stopped, NO levels rose slowly over 5 min; upon resuming US, NO release rapidly increased again. In contrast, negligible NO release occurred without US exposure, confirming US‐triggerable cleavage of the S─N bond in PMS. The finding that mSNO is stable at 37°C in vitro further underscores the essential role of US as the triggering mechanism (Figure S1j). Met release was quantified using UV–vis spectrophotometry based on a Met standard curve (Figure S1l). Release studies were conducted under lysosomal‐mimicking (pH 5.0) and physiological (pH 7.4) conditions, with or without added esterase. Maximum Met release occurred at pH 5.0 in the presence of esterase (Figure 2o), primarily due to enzymatic hydrolysis of the C─C bond linking Met to PVBC [29]. The lowest cumulative release (27.36 ± 1.68%) was observed at pH 7.4 without esterase, suggesting that under physiological conditions, release of PMet is inhibited due to partial deprotonation of amino or biguanide groups in the PMet [37]. Finally, PMS exhibited excellent hemocompatibility with hemolysis rates below 5.0% (Figure S1m).

2.2
Evaluation of Cytotoxicity of PMS in vitro
First, SK‐Hep1 cells were exposed to US irradiation at varying power levels to determine the optimal parameters that would minimize unwanted cytotoxicity of US during combined PMS and US administration. As shown in Figure S2a,b, US administration induced a power‐dependent cytotoxic effect with ∼22.15 ± 3.19% of dead cells in culture exposed to 2 W cm−2. Further, varying US powers were also tested for their ability to trigger NO release from mSNO. It was revealed that in the absence of US exposure, PMS remained stable with only a very low NO amount released to the culture media. Under low‐intensity US irradiation (0.5 W cm−2), NO started to be released from PMS, and the release increased significantly with both higher US power and PMS concentration (Figure 3a). Based on these results, the following US parameters were selected for subsequent in vitro experiments: 1 W cm−2, 1 MHz, and a 50% duty cycle, 2 min. Next, PMS and PMet were incubated with SK‐Hep1 cells and LX‐2 cells for 12 or 24 h, followed by Cell Counting Kit‐8 (CCK‐8) assays to examine their cytotoxicity. After 24 h of incubation with PMS, the viability of SK‐Hep1 cells without US irradiation remained above 84.84 ± 2.72% (at a concentration of 200 µg mL−1), while at the same concentration, the US‐irradiated cells exhibited a viability reduced to 66.81 ± 6.90% (Figure 3b). Noteworthy, viability of US‐irradiated SK‐Hep1 cells was markedly lower than viability of the non‐US cells, evidencing that US was able to ultimately trigger an efficient release of NO in a short time period [38].

2.3
Efficacy of PMS in LSECs
In the fibrotic liver microenvironment, LSECs undergo capillarization, characterization by the loss of fenestrae and formation of an organized basement membrane. This process not only precedes activation of HSCs but also forms a significant barrier to drug delivery [13]. Therefore, reversing LSECs capillarization has become an emergent and promising therapeutic target for treating liver fibrosis. Thus, we further investigated on the mechanisms by which PMS restores fenestrae in LSECs (Figure 3c) and evaluated its fenestrae‐restoring activity. SK‐Hep1 cells were pretreated with lipopolysaccharide (LPS) and incubated in high‐glucose medium followed by administration of PMS and other control treatments for 24 h. ELISA revealed that compared to the model (induced, non‐treated) group (24.73 ± 0.98 pg mL−1), the sGC levels were significantly (P < 0.0001) increased in the PMS group (47.98 ± 2.64 pg mL−1) and even more in the PMS+US group (351.48 ± 11.04 pg mL−1) (Figure 3d). Correspondingly, intracellular cGMP rose markedly in these groups, with the PMS+US group showing a 26.11‐fold increase relative to the model group (Figure 3e).
To directly assess the fenestrae repair capabilities of PMS, a transwell co‐culture model was employed using murine primary LSECs (process of isolation and sorting of LSECs is shown in Figure 3f; Figure S2c). The apical chamber was seeded with LSECs isolated from healthy or fibrotic mice, while the basal chamber contained non‐activated LX‐2 cells (Figure 3g). For experiments, PMet, Met, and PMS were labeled with fluorescein isothiocyanate (FITC) and added to the apical chamber. After 36 h of co‐culture, LSECs were harvested from the apical chamber, and fenestrae (porosity) was analyzed by scanning electron microscopy (SEM). As shown in Figure 3h, PMS+US treatment significantly restored LSEC fenestrae and sieve plates compared to other treatments. Further semiquantitative analysis revealed an increase in porosity from 0.56 ± 0.12% (model group) to 6.00 ± 0.74% (PMS+US) (Figure 3i). In the same transwell setup, we also assessed the penetration of PMet through LSECs followed by uptake in LX‐2 cells in the basal chamber. The PMS+US group displayed the strongest green fluorescence signal (Figure 3j), indicating enhanced drug transport through repaired fenestrae. These findings suggest that PMS, when activated by US to release NO, effectively restores LSECs fenestrae, facilitating drug entry into the space of Disse, followed by enhanced uptake of PMet by HSCs.

2.4
Cellular Uptake of PMS by HSCs and Evaluation of Biological Activity
Next, we explored the impact of PMet in PMS on HSCs in terms of uptake, molecular, and phenotypic response. First, the biocompatibility of PMet, a critical component of PMS able to suppress HSCs activation, was confirmed, as no significant cytotoxicity was observed in LX‐2 cells (Figure S2d). To further study cellular uptake, time‐dependent confocal laser scanning microscopy (CLSM) analysis was performed using FITC‐labeled PMet. It was found that PMet signal (green) increased within cells and co‐localized with lysosomes (red) at 4 h, confirming lysosomal uptake. Besides, we noted that the green fluorescence intensity at 6 h was largely comparable to that at 4 h, and at 6 h, partial signal redistribution from lysosomal to non‑lysosomal regions could already be observed. By 8 h, PMet gradually escapes lysosomes, and its separated signal was observable in the intracellular region outside lysosomes, which is its desired region of action (Figure 4a).
Given the promising fenestrae‐restoring effects of PMS, we again utilized a transwell system to mimic sequential PMS delivery in vivo and further examined its effect on HSCs upon LSECs penetration (Figure 4b). To understand the biological activity of PMS, expression of α‐smooth muscle actin (α‐SMA), a key marker of HSCs activation was examined. As shown in Figure 4c and Figure S2e, α‐SMA expression was significantly elevated in the model group (79.05 ± 0.96 a.u., 1.35‐fold higher than in the control group), indicating successful activation of HSCs. In contrast, the PMS+US group (61.82 ± 1.66 a.u.) exhibited α‐SMA expression levels comparable to the control group. Notably, α‐SMA expression in the PMS+US group was significantly lower than in the PMS group without US (69.21 ± 2.04 a.u.), highlighting the crucial contribution of US‐triggered NO release. These results suggest that restored LSECs fenestrae allowed for improved drug delivery to the underlying aHSCs, enabling more efficient therapeutic response. Importantly, the PMS+US group also outperformed the Met group in reducing the α‐SMA expression, underpinning the importance of the combinatorial therapeutic approach.
PMet internalized by LX‐2 cells undergoes esterases‐mediated hydrolysis, releasing Met intracellularly. This phenomenon inhibits aHSCs proliferation and migration, thereby contributing to anti‐fibrotic effects (Figure 4d) [39]. Western blot revealed that PMet, PMS, PMS+US, and Met treatments increased phosphorylation of AMPK compared to the model group (Figure 4e,f), implicating PMet as a key regulator of the AMPK signaling pathway. AMPK activation consequently suppressed HSCs proliferation by inhibiting mTOR phosphorylation (Figure 4e,g). Compared with the model group (p‐mTOR/mTOR, 1.96 ± 0.20), the PMS+US group exhibited significantly reduced p‐mTOR/mTOR levels (0.72 ± 0.18). Additionally, phosphorylation of p70S6K, which is a downstream marker of mTOR activity, was markedly decreased in the PMS+US group (Figure S2f,g), suggesting the ability of PMet to inhibit aHSCs.
Finally, a wound‐healing assay was conducted to assess aHSCs migration. After 24 h of LPS stimulation, the migratory capacity of model group cells reached 24.40 ± 5.99%. Treatment with PMet, PMS, PMS+US, and Met reduced aHSCs migration to 16.57 ± 6.57%, 14.73 ± 3.37%, 4.87 ± 3.24%, and 15.41 ± 0.94%, respectively, with the PMS+US demonstrating the most efficient inhibitory activity. In summary, we validated that PMet is a crucial part of PMS responsible for inhibition of proliferation and migration of aHSCs via the AMPK‐ mTOR‐ p70S6K signaling axis.

2.5
Biodistribution and Cellular Specificity of PMS in vivo
For the biodistribution study, ICR mice were intraperitoneally injected with either olive oil (control) or carbon tetrachloride (CCl4) for 6 weeks to establish healthy and liver fibrosis model (Figure S3a). After establishing models, 1,1‐dioctadecyl‐3,3,3,3‐tetramethylindotricarbocyanine iodide (DiR)‐labeled PMS were administered via tail vein injection. in vivo fluorescence imaging was performed at 0 h, 5 h, and on days 1, 2, 4 and 7 post‐injection to assess biodistribution in both normal and fibrotic mice. As shown in Figure S3b, peak fluorescence in the liver was observed in all groups at 2 days post‐injection. DiR‐labeled PMS demonstrated sustained bioaccumulation in the liver for up to 7 days, likely due to sequestration by the hepatic reticuloendothelial system. Notably, in healthy mice, the fluorescence declined rapidly, potentially reflecting efficient hepatic clearance. In contrast, the fibrotic group exhibited prolonged fluorescence retention, which may be attributed to impaired liver function and reduced clearance capacity caused by fibrosis.
Hepatic bioaccumulation was further analyzed via ex vivo fluorescence imaging of major organs (Figure S3c). On day 2 in the PMS+US‐treated fibrotic mice, liver fluorescence intensity was 4.91‐fold and 8.48‐fold higher than that of the spleen and lungs, respectively. Across all time point, liver bioaccumulation significantly exceeded that in other organs in both healthy and fibrotic mice (Figure S3d–f). Moreover, the fluorescence intensity in fibrotic livers was consistently stronger than in healthy livers, further supporting enhanced bioaccumulation of DiR‐labeled PMS in fibrotic livers. Notably, the liver fluorescence intensity in the PMS+US group was significantly higher than that in the PMS group, approximately 1.24‐fold of the PMS group in 2 d, which directly demonstrating that US exposure is a necessary factor for markedly enhancing hepatic accumulation of PMS under identical fibrotic model conditions. This finding strongly corroborates that the US‐induced fenestrae restoration mechanism effectively improves the ability of the nanoassemblies to traverse the liver sinusoidal endothelial barrier.
To identify the primary cell types involved in PMS uptake, fibrotic mice were intravenously injected with DiI‐labeled PMS for subsequent liver immunofluorescence analyses. CD31 and α‐SMA were used as markers for LSECs and aHSCs, respectively. As shown in Figure S3g, minimal colocalization of DiI (red) with CD31 (green) was observed, indicating limited uptake of PMS by LSECs. In contrast, significant colocalization of 1,1'‐dioctadecyl‐3,3,3',3'‐tetramethylindocarbocyanine perchlorate (DiI) with α‐SMA was detected, particularly in the PMS+US group, where a strong yellow signal (indicative of red/green overlap) was observed (Figure S3h). Together, these results confirmed importance of US allowing NO release to restore fenestrae in vivo, further enhancing PMS uptake of aHSCs.

2.6
Anti‐Fibrotic Activity of PMS in CCl4‐Induced Mice
Following confirmation of the intracellular distribution of PMS, we proceeded to evaluate its therapeutic anti‐fibrotic efficacy. Different formulations were administered via tail vein injection twice weekly for 4 weeks, and therapeutic outcomes were assessed after the final dose (Figure 5a). To comprehensively monitor improvements of liver pathological conditions, US imaging was performed (Figure 5b) [40]. The hepatorenal grayscale ratio, a diagnostic metric reflecting liver echogenicity, showed that PMS+US treatment (0.60 ± 0.03) significantly restored liver echogenicity toward physiological levels (0.53 ± 0.07) (Figure 5c). Gross morphological examination revealed smooth surfaces, soft texture, bright red color, and sharp edges in livers from healthy mice. In contrast, fibrotic mice exhibited coarse granular liver surfaces, firm texture, dull coloration, and rounded edges. All treatment groups showed varying degrees of morphological recovery, with the PMS+US group demonstrating the most pronounced improvements (Figure 5d). SEM images of liver sinusoids revealed intact fenestrae and the absence of a basement membrane in normal mice. In contrast, CCl4‐induced fibrosis led to a marked reduction or complete loss of fenestrae. Treatment with PMet, PMS, PMS+US, or Met all restored fenestrae to varying extents, with the PMS+US group exhibiting the most substantial fenestrae recovery. No significant difference was observed between the US only and CCl4‐induced group, which clearly demonstrates that the physical effects of US alone (including cavitation and mechanical forces) are insufficient to reverse LSECs capillarization. Instead, the specific recovery of fenestrae structures requires the co‐presence of both US and PMS (Figure 5e). Semiquantitative analysis confirmed that sinusoidal porosity in the PMS+US group reached 21.30 ± 1.14%, closely approximating that of healthy controls (Figure 5f). In the next step, biomarkers of liver injury were quantified. Compared with the fibrotic group, serum aspartate aminotransferase (AST) and alanine aminotransferase (ALT) were reduced across all treatment groups, with PMS+US treatment achieving the most substantial reductions (Figure 5g). Given that liver fibrosis progression involves excessive ECM deposition, hydroxyproline content was quantified as a marker of collagen accumulation [14]. PMS+US treatment significantly reduced hydroxyproline levels corroborating its efficient anti‐fibrotic activity (Figure 5h).
To explore mechanistic pathways, we quantified sGC, PKG (Figure S2h,i), and cGMP (Figure 5i) in murine hepatic tissues. Mice treated with PMS+US exhibited a 1.24‐fold increase in hepatic cGMP compared to untreated fibrotic mice. Similarly, sGC and PKG levels were significantly increased after PMS+US treatment. This suggests that PMS+US administration preserved LSECs differentiation via activation of the NO‐sGC‐cGMP pathway. Previous studies have shown that Met inhibits the proliferation and migration of aHSCs by activating AMPK and suppressing the mTOR signaling pathway and its downstream targets [41].
To validate the combined anti‐fibrotic effects of PMS and US and observed inhibitory effects of PMet on aHSCs, we conducted histological and immunohistochemical assessment of liver tissues using hematoxylin and eosin (H&E), Sirius Red staining, and Masson's trichrome staining and immunostaining for α‐SMA and collagen I. H&E‐stained liver sections from the fibrotic group showed extensive lobular necrosis, loss of cellular boundaries, tissue proliferation, and inflammatory infiltration. These pathological features were notably alleviated in all treatment groups, with PMS+US showing the largest extent of reduction of necrosis and lymphocytes infiltration (Figure S4a). Sirius Red staining revealed a 76.32% reduction in collagen deposition in the PMS+US‐ group compared to untreated fibrotic mice (Figure 5j), consistent with results from Masson's trichrome staining and collagen I immunostaining (Figure 5k; Figure S4b,c). Moreover, the α‐SMA‐positive area, indicative of aHSCs, was reduced from 10.27 ± 0.73% in the fibrotic mice to 4.92 ± 0.31% in the PMS+US group (Figure 5l). Collectively, these findings clearly demonstrate that PMS, particularly when combined with US, exhibits potent anti‐fibrotic activity through two crucial mechanisms: i) restoration of LSECs fenestrae and ii) inhibition of aHSCs via the AMPK‐mTOR signaling.

2.7
Elucidating Molecular Mechanisms of PMS Anti‐Fibrotic Activity
To further elucidate the molecular mechanisms underlying the therapeutic effects of PMS, we performed RNA‐Seq on liver tissues from healthy, fibrotic, and PMS+US‐treated mice. Hierarchical clustering revealed a substantial number of differentially expressed genes (DEGs) in the fibrotic group relative to healthy controls (Figure S5a), underscoring a distinct transcriptional signature across the three cohorts. A total of 636 DEGs were identified between the PMS+US‐treated and untreated fibrotic (model) groups, with 291 genes uniquely deregulated by PMS+US treatment (Figure 6a). Among these, 160 genes were significantly up‐regulated, and 476 were down‐regulated, indicating a robust transcriptomic shift induced by the PMS+US exposure (Figure 6b). Unsupervised principal‐component analysis (PCA) of the identified DEGs further corroborated the distinct segregation of the three groups (Figure 6c). Consistent with these findings, hierarchical clustering and volcano plots (Figure S5b,c) demonstrated markedly different transcriptomic profiles among the groups. Subsequent bioinformatic analysis revealed that numerous genes and their associated pathways are implicated in the initiation and progression of hepatic fibrosis. To delineate the molecular pathways influenced by PMS+US, we performed Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis. As shown in Figure 6d,e, twenty pathways relevant to therapeutic efficacy were identified, including fibrosis‐associated cascades and, notably, the cGMP‐PKG signaling axis. Within this pathway, nitric oxide activates sGC, leading to increased cGMP levels and the activation of PKG‐dependent signaling. This further promotes the re‐differentiation of LSECs and degradation of the pathological basement membrane, which is precisely the biological response to the PMS+US therapy. Furthermore, multiple pathways closely linked to hepatic fibrogenesis were significantly down‐regulated, underscoring the synergistic anti‐fibrotic activity of the PMS+US administration. We also observed a significant enrichment of the AMPK signaling pathway associated with HSCs. To validate this finding, we performed Western blot analysis to assess the expression of selected proteins in hepatic tissues. Compared to the fibrotic mice, phosphorylation of AMPK was significantly increased in the PMS+US group (1.18 ± 0.01 vs 0.76 ± 0.33), confirming the efficacy of PMet, which is the key active component of PMS responsible for AMPK activation. In addition, PMS+US treatment substantially inhibited phosphorylation of mTOR and its downstream effector p70S6K [42], with a pronounced decrease in the p‐p70S6K/p70S6K ratio compared to the fibrotic mice (Figure 6f). We also mapped the DEGs to the top 30 enriched Gene Ontology (GO) categories (Figure 6g). Notably, a large proportion of these genes are involved in cell differentiation, ECM organization, and cellular response to external stimuli. Heatmap visualization revealed that PMS+US treatment promotes LSEC re‐differentiation by modulating the expression of Il7, Ccr2, Tlr2, Trp63, and Cdkn2b (Figure 6h), while simultaneously mitigating fibrosis through down‐regulation of ECM overdeposition‐associated genes such as Svep1, Col6a6, Col28a1, Col4a6, and Col5a3. Collectively, the RNA‐seq data elucidated the molecular aspects by which PMS+US exerts a potent anti‐fibrotic effect by simultaneously reversing LSECs dedifferentiation and suppressing HSCs activation.

2.8
Biosafety Evaluation of US Administration and PMS+US‐Triggered NO Release
To systematically evaluate the biosafety of the US parameters used in this study (1.0 MHz, 1.0 W cm−2, 50% duty cycle, 2 min) and the associated PMS+US‐triggered NO release, we performed real‐time temperature monitoring of the insonated area using infrared thermography. As shown in Figure S6a, only a limited temperature increase was observed throughout the irradiation. The maximum surface temperature reached Tmax = 39.9°C at 2 min and Tmax = 42.4°C at 150 s. The corresponding maximal temperature rises were ΔT = 2.8 ± 0.3°C and ΔT = 4.4 ± 0.06°C, respectively, and in both cases the temperature remained below the short‐term thermal safety threshold of 43°C [43]. These results indicated that the US settings applied here do not cause appreciable heat accumulation and are unlikely to pose a detectable risk of thermal injury. To assess potential local effects of US exposure on healthy tissues, the livers from healthy and healthy+US groups were excised for gross inspection, and no obvious hemorrhage/tissue damage was observed (Figure S6b). The insonated skin region was further examined by H&E staining (Figure S6c). The epidermis remained continuous and intact, and skin appendages such as hair follicles and sebaceous glands were clearly preserved. No necrosis, marked edema, or substantial inflammatory cell infiltration was detected, and no hemorrhagic features (e.g., erythrocyte extravasation) were observed. These histological findings further support good local tissue tolerability under our ultrasound conditions. We further performed H&E examinations of major organs, and no evident pathological alterations, such as inflammatory infiltration, hemorrhage, necrosis, or structural disruption, were observed in any group after ultrasound exposure (Figure S6d). In addition, serum biochemistry (blood urea nitrogen (BUN), aspartate transaminase (AST), alanine transaminase (ALT) and uric acid (UA) (Figure S6e), and coagulation‐related parameters fibrinogen (FIB) and activated partial thrombin time (APTT) (Figure S6f) showed no significant differences between the US and control groups, indicating that this strategy does not introduce an apparent risk of coagulation dysfunction.
Given that systemic NO exposure may cause vasodilation and hypotension, we monitored mouse blood pressure during the treatment course. As shown in Figure S6g,h, following injection in the mSNO+US group, mice exhibited a pronounced and sustained decrease in blood pressure (mean reduction in systolic blood pressure > 25 mmHg), with slow recovery during the monitoring period and values remaining below the normal range. This confirms that systemic NO release indeed produced the expected hypotensive effect. In contrast, blood pressure in the PMS group remained stable. Meanwhile, the blood‐pressure profiles of the PMS+US and US only groups were highly overlapping, both groups showed an immediate increase in blood pressure after ultrasound exposure, which we attribute to the US‐associated stress response. Importantly, blood pressure returned to normal within 2 h and remained stable thereafter. Collectively, these data provide strong evidence that, by enabling local triggering and site‐specific release, our smart nanoassembly successfully decouples the therapeutic effects of NO from its potential systemic cardiovascular side effects, thereby ensuring overall systemic safety.

2.9
Assessment of Biosafety of PMS+US Therapeutic Intervention
The biosafety profile of PMS+US therapeutic intervention was further assessed through analyzing an array of parameters in vivo. Key serum biomarkers associated with inflammation and organs function, including BUN, total cholesterol (TC), triglyceride (TG), alkaline phosphatase (ALP), urea and creatinine (CREA) were quantified to evaluate possible systemic toxicity of PMS (Figure S7a–f). No significant differences in these parameters were observed between the healthy group and any of the treated groups, indicating no apparent hepatic, renal, or metabolic toxicity. In parallel, we monitored mice body weights trends throughout the treatment period (Figure S7g). Healthy mice exhibited a gradual increase in body weight, consistent with normal growth. In contrast, fibrotic mice showed body weight loss, caused by acute hepatotoxicity of CCl4. Interestingly, the PMS+US group displayed a slightly lower average body weight post‐treatment, potentially reflecting mild stress or discomfort associated with repeated US exposure. Nevertheless, this did not correlate with any observable pathological damage, thus being considered safe. Histopathological examination of H&E‐stained sections from major organs, including the spleen, kidneys, lungs and heart, revealed no significant tissue abnormalities or signs of undesired organ toxicity in any treatment group (Figure S7h). Together, these findings confirm that PMS combined with US exposure exhibit a favorable biosafety profile and are well tolerated in vivo.

3
Conclusions
In summary, we have rationally designed and validated superb properties of a dual‐prodrug (mSNO and PMet) polymeric delivery system, PMS, for targeted therapy of liver fibrosis. PMS is specifically engineered to restore the fenestrated phenotype of LSECs, thereby enhancing drug delivery in the space of Disse. Through co‐culture models, we demonstrated that PMS effectively repairs capillarized LSECs, leading to increased fenestrae formation and improved permeability, which facilitates drug transport to HSCs. Simultaneously, the co‐culture experiments reduced expression of α‐SMA, a hallmark of aHSCs, indicating therapeutic efficacy. Mechanistically, under US irradiation, PMS first releases NO, which binds to Fe2
+ in heme group of sGC, activating the NO‐sGC‐cGMP signaling pathway, promoting LSECs re‐differentiation and fenestrae restoration. The remaining part of PMS, PMet, then crosses the repaired fenestrae and intercellular gaps to reach the Disse space, where it is internalized by HSCs. Within HSCs, intracellular esterases trigger the release of active Met, leading to inhibition of HSC activation, proliferation and differentiation. In both in vitro and in vivo models, PMS significantly promoted the reversion of capillarized LSECs to their native fenestrated phenotype and suppressed fibrogenic activation of HSCs. These findings highlight the therapeutic potential of PMS as a multifunctional delivery system and pave new avenues for targeted drug delivery across the sinusoidal barrier. Overall, this work introduces the “fenestrae restoration first, then therapeutic delivery” paradigm, providing a promising and effective approach for liver fibrosis by enhancing drug transport into the space of Disse and targeting key pathogenic cell populations.

4
Experimental Section
4.1
Materials and Reagents
VBC, styrene (St), 2,2'‐azobis(isobutyronitrile) (AIBN), and N,N‐diisopropylethylamine (DIPEA) were purchased from Meryer Chemical Technology Co., Ltd. (Shanghai, China). Toluene was obtained from Yuanli Chemical Co., Ltd. (Tianjin, China). Met, pyrene, FITC, tert‐butyl nitrite (TBN), and methanol were sourced from Heowns Science Co., Ltd. (Tianjin, China). Dimethyl sulfoxide (DMSO) was acquired from Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). Thiol‐terminated poly(ethylene glycol) (mPEG‐SH, MW = 2000) was purchased from Shanghai Yare Biotech Co., Ltd. (Shanghai, China). CCK‐8, Griess reagent assay kit, DiI, and LPS, along with the following antibodies were obtained from Beyotime Biotechnology. (Shanghai, China): rabbit anti‐AMPK alpha 1 monoclonal antibody (AF1627, 1:1500 dilution), rabbit anti‐phospho‐AMPKα1(Thr183)/AMPKα2(Thr172) polyclonal antibody (AF5908, 1:1000), rabbit anti‐p70S6K polyclonal antibody (AF0258, 1:600), rabbit anti‐phospho‐p70S6K (Thr389) polyclonal antibody (AF5899, 1:2000), rabbit anti‐mTOR monoclonal antibody (AF1648, 1:1000), rabbit anti‐phospho‐mTOR (S2448) polyclonal antibody (AF5869, 1:1000), rabbit anti‐α‐Smooth muscle actin polyclonal antibody (AF0048, 1:100), rabbit anti‐COL1A1/Collagen I monoclonal antibody (AF1840, 1:100), goat anti‐IgG secondary antibody conjugated with horseradish peroxidase (HRP) (A0208, 1:1000). Rabbit anti‐β‐Actin polyclonal antibody (21338, 1: 1500) was obtained from Signalway Antibody Co., Ltd. (Shanghai, China). Dulbecco's modified eagle medium (DMEM), minimum essential medium (MEM), FBS, non‐essential amino acids (NEAA), sodium pyruvate, and penicillin/streptomycin were obtained from Gibco (Grand Island, NY, USA). Roswell Park Memorial Institute 1640 (RPMI‐1640) was acquired from SenBeiJia Biological Technology Co., Ltd. (Nanjing, China). CCl4 and olive oil were purchased from Macklin Biochemical Co., Ltd. (Shanghai, China). Hoechst 33342, LysoTracker Red, Percoll, and polylysine were provided by Solarbio Science & Technology Co., Ltd. (Beijing, China). Mouse and human sGC, cGMP, and PKG ELISA kits were purchased from Meimian Industrial Co., Ltd. (Jiangsu, China). X‐RIPA UltraMix, X‐strip, X‐Loading were purchased from X‐BLOT lifeScience Ltd. (Suzhou, China). DiR was bought from Uelandy Biotechnology Co., Ltd. (Suzhou, China). Recombinant Murine EGF was bought from Thermo Fisher Scientific (Waltham, MA, USA). Esterase derived from porcine liver, ≥15 units mg−1 solid, was acquired from Yuanye Bio‐Technology Co., Ltd. (Shanghai, China). 3% glutaraldehyde was bought from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China), and osmium tetroxide was obtained from Zhongjing Keyi Technology Co., Ltd. (Beijing, China). All chemical reagents were used as purchased without further purification.

4.2
Formulation of PMS
4.2.1
Preparation of PVBC
PVBC was synthesized via RAFT polymerization. For the copolymerization of St and VBC, St (9.0 g, 90 mmol), VBC (1.17 g, 7.6 mmol), a carboxylic acid‐functionalized trithiocarbonate RAFT agent (R2) (0.365 g, 1.0 mmol), AIBN (0.052 g, 0.31 mmol), and toluene (5 mL) were added into a round‐bottomed flask. The flask was evacuated to establish a vacuum environment, then purged with N2 and immersed in an oil bath at 80°C to initiate polymerization. The reaction was carried out for 5 h under strictly anhydrous and oxygen‐free conditions to maintain the integrity of the reaction [36, 44]. Upon completion, the reaction mixture was poured into excess methanol to precipitate the polymer. The resulting solid was dried under vacuum at 40°C for 48 h. For further purification, the crude polymer was washed with a solvent system comprising petroleum ether and ethyl acetate (3:1 v/v).

4.2.2
Preparation of PMet
PVBC (34.708 mg), HCl·Met (433.5 mg), and DIPEA (679 µL) were dissolved in DMSO and stirred in an oil bath at 130°C for 72 h. The reaction mixture was dialyzed against hydrochloric acid (pH 3.5) for 3 d, and distilled water for an additional 2 d. The Met prodrug (PMet) was obtained by lyophilization.

4.2.3
Preparation of mSNO
The NO prodrug, mSNO, was synthesized by reacting the ─SH group of mPEG‐SH with TBN. Briefly, mPEG‐SH (20 mg) and an excess amount of TBN (∼20 mg) were dissolved in methanol (10 mL) and stirred at 4°C under N2 atmosphere for 24 h in light‐protected conditions. The resulting product with ─SNO groups was quickly dried under vacuum for 5 min and then stored at −20°C in the dark.

4.2.4
Formulation of PMS
To prepare the PMS nanoassembly, PMet and mSNO were bound through electrostatic interactions driven by their opposite charges. The as‐prepared PMet and mSNO were stirred in an ice bath for 4 h under light‐protected conditions. The resulting mixture was centrifuged at 8000 rpm, and the pellet was washed three times with anhydrous ethanol and deionized water, respectively. The final product was lyophilized and stored at −20°C in the dark for further use. For fluorescence labeling of PMS, FITC‐labeled PMS was prepared by stirring PMS (30 mg) with FITC (1 mg) in deionized water overnight, allowing covalent attachment of the fluorophore for subsequent tracing experiments.

4.3
Physico‐Chemical Characterization of PMS
The morphology and size analyses were carried out using TEM (Tecnai G2 F20, FEI, Eindhoven, Netherlands). Hydrodynamic diameter and zeta potential were measured via DLS and ELS using ZetaSizer (Nano ZS, Malvern Instruments, Malvern, UK). 1H NMR (Ascend 400, Bruker) spectra were recorded at 25°C in deuterated chloroform (CDCl3) as the solvent. UV–vis absorption spectra were recorded using a UV–vis spectrophotometer (Cary 60, Agilent, USA). FT‐IR spectra were recorded using an infrared spectrometer (TENSOR 27, Bruker, Bremen, Germany). XPS analyses were performed using an XPS spectrometer (ESCALAB‐250Xi, Thermo Fisher Scientific). The molecular weight of polymers was analyzed using a gel permeation chromatography (GPC, Waters 1515, Shanghai, China).

4.4
Determination of CMC
A pyrene stock solution and a series of PMet solutions at different concentrations were prepared. The pyrene solution was added into brown Eppendorf tubes and allowed to dry to remove acetone. Subsequently, micelle solutions of varying concentrations were added. The mixtures were shaken at 65°C for 2 h and left overnight in the dark room at room temperature. Fluorescence emission intensities at 384 nm and 373 nm were recorded, and the fluorescence intensity ratio (I384/I373) was plotted against the logarithm of the sample concentration (Log C). The CMC value of PMet was determined by identifying the intersection point of the two fitted linear regions.

4.5
Evaluation of in vitro Release Kinetics of NO and Met from PMS
NO release from PMS was quantified using a Griess reagent, which detects nitrite ions formed from NO oxidation. PMS solution (2 mg mL−1, 4 mL) was treated with an ultrasonic probe (1 W cm−2) in the dark. At predetermined time points, 50 µL aliquots were withdrawn and added to transparent 96‐well microplates. After adding Griess reagent, absorbance at 540 nm was measured using a microplate reader (Infinite 200 PRO, Tecan, Männedorf, Switzerland). NO concentration was calculated from a standard curve of NaNO2. Met release from PMet was examined via a dynamic dialysis. PMet was placed in a dialysis bag (MWCO 3.5 kDa) and immersed in phosphate‐buffered saline (PBS, pH 7.4 and pH 5.0) at 37°C with stirring at 120 rpm. At set intervals, the medium was removed and replaced with an equal volume of fresh PBS. Released Met was quantified using a Nanodrop spectrophotometer (NanoDrop One/OneC, Thermo Fisher Scientific), with concentrations calculated from a standard curve.

4.6
Evaluation of Hemolytic Properties of PMS
Whole blood was collected from healthy ICR mice and centrifuged at 3000 rpm for 15 min to collect erythrocytes. Erythrocytes were washed 3–4 times with PBS and diluted tenfold. A volume of 200 µL of erythrocyte suspension was mixed with PMS (1 mL) at various concentrations and incubated at 37°C for 4 h. After centrifugation at 12 000 rpm for 15 min, the absorbance of the supernatant was recorded at 570 nm. PBS and deionized water served as negative and positive controls, respectively. The hemolysis rate (%) was calculated using the formula:

4.7
Cell Culture Conditions
The human HSC line LX‐2 was sourced from Procell (Wuhan, China). The human LSEC cell line SK‐Hep1 was a kind gift from Associate Professor Meng Yu (Southern Medical University, Guangzhou, China). LX‐2 cells were cultured in DMEM with 10% FBS and 1% penicillin‐streptomycin. SK‐Hep1 cells were maintained in MEM with 10% FBS, 1% NEAA, 1% sodium pyruvate, and 1% penicillin‐streptomycin. All cell cultures were cultured at 37°C in a humidified atmosphere containing 5% CO2.

4.8
Optimization of Ultrasound Intensity and Evaluation of Cytotoxicity of PMS
We used Nu‐Tek therapeutic ultrasound system UT1021 (Sonic‐Stimu Basic) to determine the optimal ultrasonic intensity for inducing NO release while minimizing undesired cytotoxicity, SK‐Hep1 cells were treated with PMS and exposed to various ultrasonic powers. Cell viability was assessed via Calcein‐AM/PI staining. The NO released under different PMS concentrations and ultrasound intensities was also quantified. Cytotoxicity was evaluated using the CCK‐8 kit. SK‐Hep1 and LX‐2 cells were seeded into 96‐well plates (∼8 × 103 cells/well) and incubated for 24 h. Cells were then treated with varying concentrations of PMS and PMet (0, 6.25, 12.5, 25, 100, 200 µg mL−1) for 12 or 24 h. After treatment, the medium was removed, cells were washed with PBS, and CCK‐8 reagent (0.1 M) was added. After incubation at 37°C for 2 h, absorbance at 450 nm was measured.

4.9
Isolation of LSECs
Livers were collected from healthy or fibrotic mice, washed with PBS (containing 1% penicillin‐streptomycin) and transferred to a dish with digestion buffer. After removal of the connective tissues, livers were minced and grounded using a sterile syringe. The suspension was transferred to centrifuge tubes and incubated at 37°C for 15 min with shaking. Following digestion, the mixture was filtered through a 200‐mesh cell strainer and centrifuged at 100×g for 5 min. The supernatant was collected, resuspended in RPMI‐1640, and centrifuged again. Supernatants from both centrifugations were combined and further centrifuged at 400×g for 10 min. The pellets were resuspended in 25% Percoll and carefully layered over 50% Percoll to form a distinctive interface. After centrifugation 900×g for 20 min, the LSEC‐enriched interface between 25% and 50% Percoll was collected, diluted tenfold with RPMI‐1640, and centrifuged at 2000×g for 10 min. Cells were seeded onto poly‐L‐lysine‐coated coverslips in 12‐well plates. containing RPMI‐1640 with 15% FBS and 10 ng mL−1 recombinant murine EGF. Cells were incubated for 24–48 h before use.

4.10
Cellular Uptake
LX‐2 cells were seeded into confocal dishes (∼5 × 105 cells/dish) and treated with FITC‐labeled PMet (100 µg mL−1) for 0, 2, 4, 6, and 8 h. After washing, LysoTracker Red and Hoechst 33342 were used to stain lysosomes and nuclei, respectively. CLSM (Leica, Wetzlar, Germany) was performed using the following excitation/emission settings: Hoechst 33342: 405/491 nm; LysoTracker Red: 577/590 nm. Colocalization was analyzed using Image J (National Institute of Health, Bethesda, MA, USA).

4.11
Immunofluorescence
A transwell system was employed to investigate how LSEC fenestrae influence drug uptake in HSCs. Primary murine LSECs and LX‐2 cells were seeded in the apical and basal chambers, respectively. After cells attachment, FITC‐labeled PMS was added to the apical chamber and incubated for 24 h. Nuclei were stained with Hoechst 33342 for 20 min, and PMet internalized by HSCs was imaged by CLSM. To evaluate activation of markers of aHSCs, cells were incubated with α‐SMA primary antibody overnight at 4°C. After washing with PBS, FITC‐conjugated secondary antibody was added, and cells were incubated at room temperature for 1 h. Hoechst 33342 was used for nuclear staining. CLSM images were analyzed using Image J for quantification.

4.12
SEM of LSECs
24 h post‐treatment, murine primary LSECs from the apical chamber were collected and seeded onto coverslips. After attachment, the cells were fixed in 3% glutaraldehyde. Then, cells were washed three times with ultrapure water, coated with 1% osmium tetroxide, and washed three times with ultrapure water. Dehydration was carried out with ethanol gradient (30%→50%→70%→90%→100% ×3), 15 min each. Then, samples were placed into a critical point dryer for drying followed by gold sputtering. SEM imaging was performed using JEOL JSM‐IT700HR (JEOL, Tokyo, Japan), and liver sinusoidal porosity was quantitatively analyzed using Image J.

4.13
Western Blotting
For Western blotting, total proteins were extracted using X‐RIPA UltraMix buffer supplemented with phosphatase inhibitors cocktail. Before separation, total proteins were quantified using the BCA assay. 30 µg of proteins was separated using sodium dodecyl sulfate‐polyacrylamide gel electrophoresis (SDS‐PAGE) followed by transfer onto polyvinylidene fluoride membranes, subsequently blocked with blocking solution for 10 min. Membranes were washed three times with Tris buffered saline with Tween‐20 for 15 min. After blocking, primary antibodies were added in dilutions described in Materials and Reagents section and membranes were incubated at 4°C overnight. After washing, membranes were incubated with HRP‐conjugated secondary antibodies for 2 h at room temperature, and membranes were visualized using ECL reagent and the Bio‐Rad ChemiDoc MP system (Bio‐Rad, Hercules, CA, USA). Protein expression was densitometrically quantified using Image J. It should be noted that the total protein was re‐probed after stripping the membrane used for phospho‐protein detection, followed by re‐incubation and subsequent procedures.

4.14
Examination of Effect of PMS on HSCs Migration
A wound‐healing assay was conducted to evaluate the inhibitory effect of PMS on HSCs migration. When LX‐2 cells confluence exceeded 95%, a sterile 200‐µL pipette tip was used to create a vertical scratch across the cell monolayer. After three PBS washes, the medium was replaced with serum‐free medium with or without the annotated treatments, and this time point was recorded as 0 h. Images were captured using an inverted microscope (TS2, Nikon, Tokyo, Japan) until the experimental endpoint (24 h).

4.15
Establishment of Murine Experimental Model of Liver Fibrosis
Male ICR mice (8 weeks old) were obtained from Huafukang Biological Technology Co., Ltd. (Beijing, China). All animals were housed under controlled temperature and humidity conditions, and all procedures followed institutional guidelines approved by the Animal Ethics Committee of Tianjin University (approval no. SYXK‐2019‐0002), in accordance with the Declaration of Helsinki. Liver fibrosis was induced via intraperitoneal injection of CCl4 diluted in olive oil (1: 9) twice weekly for 6 weeks. A healthy control group received equal intraperitoneal injections of olive oil. Since Met is a component of PMS forming PMS with mSNO at a 1:1 mass ratio, a total PMS dose of 10 mg kg−1 corresponds to a PMet dose of 5 mg kg−1. Based on our measured Met loading in PMet (∼2.43%), the actual Met‐equivalent dose is 0.12 mg kg−1.

4.16
in vivo Safety Evaluation of US‐Triggered NO Release
To systematically evaluate the biosafety of the US parameters used in this study, we performed real‐time temperature monitoring of the insonated area using a handheld infrared thermal imaging camera (Hikvision UD18354B) in an in vitro phantom setup (PMS dispersed in a 37°C water bath). We also analyzed that NO release was strictly confined to the local hepatic region exposed to US, thereby preventing systemic diffusion and potential cardiovascular side effects such as hypotension. Fibrotic mice were randomly assigned to four groups: PMS, US only, PMS+US, and free NO donor (mSNO+US). Blood pressure was systematically monitored throughout the treatment period using a tail‐cuff system Medlab‐1DX single‐channel non‐invasive blood pressure measurement device (Nanjing Calvin Biotechnology Co., Ltd.). All animals underwent a 5‐day acclimation training before measurements. Systolic blood pressure (SBP) and diastolic blood pressure (DBP) were recorded continuously at 0 h (baseline) and 1, 2, 4, and 6 h post‐injection.

4.17
in vivo and Ex Vivo Biodistribution Studies
To assess tissue biodistribution, DiR‐labeled PMS was intravenously injected into healthy and fibrotic mice (including PMS‐treated and PMS+US‐treated, n = 3 per group). The comparison between the groups with and without US was done to determine how US exposure affects hepatic accumulation of PMS under the same fibrotic model conditions. At designated time points (0 h, 5 h, day 1, 2, 4, and 7), in vivo fluorescence images were captured using an in vivo imaging system (Tanon ABL‐X5; Shanghai Tanon, China) under isoflurane anesthesia. After imaging, mice were sacrificed, and major organs (liver, heart, spleen, lungs, and kidneys) were harvested for ex vivo imaging. Fluorescence intensity in each organ of the mice was quantified for statistical analysis.

4.18
Tissue Immunoanalyses and Histology
To investigate LSEC fenestrae restoration and drug uptake after US administration, fibrotic mice were intravenously injected with DiI‐labeled PMet or PMs daily for 3 days. 24 h after the final injection, liver tissues were harvested and cryopreserved for immunostaining. α‐SMA was used as marker of aHSCs. The degree of colocalization between the aHSCs and DiI‐labeled drugs was evaluated. In addition, liver tissues from different groups were fixed in 4% paraformaldehyde, dehydrated in ethanol, and embedded in paraffin. Tissue sections were stained with Sirius Red, Masson's Trichrome, hematoxylin, and eosin (H&E), and antibodies against collagen I and α‐SMA. Image J was used for quantitative analyses of stained tissues.

4.19
SEM of Liver Sinusoids
Two days after the final drug administration, mice were anesthetized. The abdominal cavity was opened to expose the liver. The portal vein and inferior vena cava were exposed. The superior vena cava was clamped with a vascular clamp to obstruct the blood flow. Subsequently, the portal vein was clamped with an arterial clamp, followed by transection of the portal vein. Needle of the perfusion apparatus was slowly inserted into the inferior vena cava, the arterial clamp on the portal vein was released, and the perfusion was initiated. When the liver lobe began enlarged, the inferior vena cava was incised, and the perfusion continued for 10 min. Liver tissue was then excised and cut into 3 × 3 × 3 mm cubes. Sample fixation, dehydration, drying, and gold sputtering for SEM analyses were performed as described above. SEM imaging was performed, and liver sinusoidal porosity was quantitatively analyzed using Image J.

4.20
Evaluation of Anti‐Fibrotic Effects In Vivo
Mice were divided into seven groups: G1: Healthy (no CCl4, olive oil‐administered), G2: Model (CCl4‐induced), G3: PMet, G4: PMS, G5: US, G6: PMS+US, G7: Met. 48 h after the final treatment, mice were euthanized, and major organs and blood samples were collected. Organs were stored in paraformaldehyde (5 mL) for fixation. In addition to histological analyses, biochemical analyses of BUN, TC, TG, ALP, urea, and CREA were conducted to assess anti‐fibrotic efficacy of PMS.

4.21
RNA‐Seq by Next‐Generation Sequencing
Livers harvested from control (healthy), fibrotic (model), and PMS+US‐treated mice were used for RNA extraction and subsequent transcriptome profiling. Total RNA was isolated with TRIzol reagent and paired‐end libraries were constructed following the manufacturer's instructions of the ABclonal mRNA‐seq Library Prep Kit (ABclonal, China). Sequencing was performed on an Illumina NovaSeq 6000 or MGISEQ‐T7 platform. Hierarchical clustering was applied to visualize gene‐expression patterns among the three groups. DEGs were subjected to GO and KEGG analyses to delineate the associated biological functions and signaling pathways. For DEGs identified between the PMS+US and model groups, expression matrices were extracted, and heat‐map analysis was conducted to compare transcript abundance across individual samples.

4.22
Liver US Imaging
Prior to imaging, the ventral area of each mouse (n = 3 per group) was depilated. After 4 weeks of treatment, liver and kidney imaging was performed using the VINNO6 Lab‐X10‐23L ultrasound imaging system. Grayscale intensity ratios between liver and kidney tissues were calculated.

4.23
Statistical Analysis
All data are presented as mean ± standard deviation (SD). Comparisons between two groups were performed using an unpaired Student's t‐test. For multiple comparisons, one‐way ANOVA with Bonferroni correction was applied. GraphPad Prism 10 (San Diego, CA, USA), was used for all statistical analyses. A p‐value of *
p < 0.05, **
p < 0.01, and ***
p < 0.001 was considered statistically significant.

Author Contributions
N.L. supervised and conceptualized the work and participated in data interpretation. S.T.L. synthesized the materials, completed the in vitro and in vivo experiments, and wrote the original draft. Z.H. and S.K. assisted with writing the original draft, critical revising the manuscript, and participated in data interpretation. M.Y.Z., X.Y.L., and Z.Y.L. participated in the in vitro and in vivo experiments and data acquisition. Z.T.Q. and Y.S.D. provided advice for the cell experiments and participated in designing experiments. All authors approved the final version of the manuscript.

Funding
This work was supported by the National Natural Science Foundation of China (W2512103, 82273873), the Tianjin Youth Science and Technology Talent Program (QN20230215), and the Tianjin Natural Science Foundation (24JCZDJC01110).

Conflicts of Interest
The authors declare no conflict of interests.

Supporting information

Supporting File: adma72864‐sup‐0001‐SuppMat.docx.