Dual‐Targeted Nanotherapy Restores Redox Homeostasis and Suppresses Uterine Hypercontractility for Effective Preterm Birth Intervention

ABSTRACT

Preterm birth (PTB), defined as delivery between 28 and 37 weeks of gestation, is a leading cause of global neonatal mortality. Its pathogenesis is primarily driven by oxidative stress and inflammation, synergistically inducing calcium ion influx into uterine smooth muscle cells, triggering aberrant contractions and PTB. Current therapies primarily offer only symptom suppression without addressing the underlying etiology, highlighting an urgent need for targeted interventions. Herein, we develop TPT, a multi‐bioactive, amphiphilic conjugate, which is synthesized through stepwise covalent conjugation of hydrophilic polyethylene glycol, a superoxide dismutase mimetic, and a hydrogen peroxide‐scavenging/anti‐inflammatory generating unit onto a molecular skeleton. TPT can self‐assemble into a multifunctional nanotherapy (designated as TPT NP). In both in vitro and in vivo lipopolysaccharide‐induced PTB models, TPT NP treatment significantly mitigates oxidative/inflammatory cascades, reduces calcium influx and apoptosis in uterine smooth muscle cells, and suppresses myometrial contractions, thereby effectively delaying PTB. Mechanistically, TPT NP restores redox homeostasis in lipopolysaccharide‐induced PTB by reducing oxidative damage products and bolstering endogenous antioxidant defenses, while concurrently improving uteroplacental hemodynamics and attenuating uterine hypercontractility. Critically, in vivo evaluations demonstrate excellent safety profiles of TPT NP, with no adverse effects on maternal health and offspring development, underscoring its significant clinical translational potential.

A newly engineered multifunctional nanotherapy, formed through self‐assembly of a multi‐bioactive amphiphilic conjugate, significantly delays preterm birth in mice. This is mechanistically achieved by mitigating oxidative/inflammatory cascades, reducing calcium influx and apoptosis in uterine smooth muscle cells, and suppressing myometrial contractions. Critically, this nanotherapy demonstrates no adverse effects on maternal health or offspring development, highlighting its significant translational promise.

1. Introduction

Preterm birth (PTB) affects 5%–18% of pregnancies worldwide and constitutes the leading cause of infant mortality and morbidity [1, 2]. It is also recognized as a pivotal risk factor for long‐term pediatric complications involving neurological, respiratory, and metabolic systems [3, 4, 5]. The pathogenesis of PTB arises from a complex interplay of maternal, fetal, and environmental factors. Among these, intrauterine infection and inflammation represent the most prevalent triggers, with subsequent oxidative stress acting as a central pathological process that disrupts gestational homeostasis [6, 7]. This core mechanism, along with other maternal contributors (e.g., gestational hypertension, diabetes, cervical insufficiency), fetal factors (e.g., multiple gestation, growth restriction), and environmental influences (e.g., substance use, chronic stress), collectively drives preterm labor [8, 9, 10, 11, 12, 13, 14].

PTB can impair fetal organ maturation, predisposing newborns to acute complications including respiratory distress syndrome, infections, hypoglycemia, and intracranial hemorrhage, while concurrently increasing maternal risks of adverse outcomes such as postpartum hemorrhage and puerperal infection. This dual threat considerably endangers both maternal and infant health [15, 16]. Current clinical management of PTB primarily employs uterine contraction inhibitors (e.g., atosiban, ritodrine), which delay labor progression by suppressing myometrial contractility [17, 18]. Concurrent administration of glucocorticoids like dexamethasone may accelerate fetal lung maturation, thereby reducing the incidence of neonatal respiratory distress syndrome [19]. In cases of cervical insufficiency, cervical cerclage serves as an effective surgical intervention to maintain cervical integrity and prevent premature dilation [20, 21]. Nevertheless, these strategies predominantly alleviate the symptomatic manifestations of PTB rather than targeting its underlying pathological mechanisms, particularly oxidative stress and inflammatory activation [22, 23]. Under physiological conditions, uterine homeostasis is maintained through a delicate balance between redox equilibrium and controlled inflammatory signaling [24]. However, risk factors can disrupt this balance, inducing excessive oxidative stress that compromises placental vascular function, amplifies inflammatory responses, and ultimately triggers uterine contractions and cervical remodeling [25]. Existing therapies exhibit limited efficacy in mitigating these core pathological processes [26]. When oxidative and inflammatory insults exceed endogenous compensatory capacity, PTB ensues [27]. Consequently, a therapeutic strategy capable of simultaneously modulating oxidative stress, inflammation, and uterine contractility holds significant promise for the effective treatment of PTB.

Recently, nanotherapies have emerged as a promising strategy for treating diverse gynecological and pregnancy‐associated diseases, including endometriosis [28], vaginal infections [29, 30], pregnancy‐related deep vein thrombosis [31], intrauterine growth restriction [32, 33], uterine scarring [34], pre‐eclampsia [35], PTB [36], and choriocarcinoma [37]. For PTB treatment, nanotherapies are designed to enhance uterine targeting and mitigate side effects of loaded drugs [38, 39]. However, a fundamental limitation persists: most current nanotherapies rely on loading traditional pregnancy‐sustaining agents like progesterone [40]. While nanocarrier engineering is sophisticated, the therapeutic efficacy remains constrained by drugs that cannot adequately address the core pathological drivers of PTB, that is, oxidative stress and inflammation [41, 42]. This disconnect explains the limited success in preventing infection‐ or inflammation‐induced PTB and underscores the critical need to develop novel therapeutics that integrate targeted delivery with intrinsic antioxidant and anti‐inflammatory activities.

Herein, we report the development of an amphiphilic multi‐bioactive macromolecular conjugate, that is, TPT, engineered through the sequential, precise covalent conjugation of cyanuric chloride (TCT), polyethylene glycol (PEG), a superoxide dismutase (SOD)‐mimetic module, and a hydrogen peroxide (H₂O₂)‐scavenging/bioactive compound‐generating moiety (Figure 1a). This conjugate self‐assembles into a multifunctional nanotherapy (designated TPT NP) designed to target the pathophysiology of PTB in a murine model. In vitro, TPT NP exhibited potent intrinsic antioxidant and anti‐inflammatory activities in uterine smooth muscle cells (USMCs). Crucially, this nanotherapy effectively attenuated calcium overload in USMCs, thereby reducing abnormal contractions and apoptosis. In vivo, TPT NP markedly mitigated oxidative‐inflammatory responses at the maternal–fetal interface, preventing PTB (Figure 1b). Importantly, TPT NP demonstrated excellent biocompatibility, with no detectable toxicity to either dams or fetuses. These findings highlight the potential of TPT NP as a safe and effective nanotherapeutic alternative to conventional tocolytics.

FIGURE 1.

Schematic illustration of the engineering of a multifunctional nanotherapy TPT NP for the treatment of PTB. (a) Development of a nanotherapy with multiple bioactivities by self‐assembly of a multifunctional amphiphilic conjugate TPT. The bioactive conjugate TPT is synthesized by covalent conjugation of TCT with three functional modules, including PEG, Tempol, and 4‐(hydroxymethyl)phenylboronic acid pinacol ester (PBAP), which can spontaneously self‐assemble into micelle‐like NPs, designated as TPT NP. (b) A sketch showing targeted treatment of PTB mice by TPT NP.

2. Results

2.1. Design and Engineering of a Bioactive Conjugate for the Treatment of PTB

Given the pathogenesis of PTB, we hypothesize that therapies capable of simultaneously inhibiting oxidative damage and pro‐inflammatory responses across multiple pathologically relevant cells represent promising therapeutic strategies. To validate this concept, we designed TPT, an amphiphilic, multi‐bioactive macromolecular conjugate. TPT features a TCT core scaffold covalently functionalized with a hydrophilic PEG segment, a SOD‐mimetic module (Tempol), and a dual‐functional moiety (phenylboronic acid pinacol ester, PBAP) capable of scavenging reactive oxygen species (ROS) and generating an anti‐inflammatory compound p‐(hydroxymethyl)phenol (HMP). Of note, PEG is well‐documented for its excellent biocompatibility, contributing to prolonged in vivo circulation of many macromolecular/nanotherapeutic agents and reduced immunogenicity [43, 44]. Concurrently, both Tempol and PBAP exhibit inherent antioxidative and anti‐inflammatory properties, enabling efficient broad‐spectrum ROS scavenging [45, 46]. Crucially, ROS‐triggered hydrolysis of PBAP generates HMP, an effective antioxidative and anti‐inflammatory phenolic compound naturally occurring in diverse plants. HMP is also a major active pharmaceutical ingredient in Gastrodia elata Blume, a widely used Chinese herb [47, 48]. These attributes provide a strong rationale for developing macromolecular therapies that integrate long‐circulation, antioxidative, and anti‐inflammatory functionalities. Furthermore, the hydrophilic PEG segment combined with the hydrophobic Tempol and PBAP, imparting amphiphilicity to the resulting conjugate. This amphiphilicity enables the self‐assembly of TPT into a nanotherapy.

TPT was synthesized through stepwise nucleophilic substitution reactions of TCT with methoxy PEG amine (mPEG‐NH₂), Tempol, and PBAP (Figure S1a). The successful synthesis of TPT was confirmed by ¹H NMR and Fourier transform infrared (FT‐IR) spectroscopy (Figure S1b,c). Observation by transmission electron microscopy (TEM) revealed a well‐defined spherical shape for the TPT‐assembled nanotherapy (TPT NP). Dynamic light scattering (DLS) measurement indicated that TPT NP displayed a narrow size distribution, with an average hydrodynamic diameter of 90 nm and a near‐neutral ζ‐potential of −2.48 ± 0.02 mV (Figure 2a,b; Figure S1d). Moreover, TPT NP was able to effectively eliminate different types of ROS, including superoxide anion, 2,2‐diphenyl‐1‐picrylhydrazyl (DPPH, a radical mimic), hydroxyl radicals, and H₂O₂, in dose‐dependent manners (Figure 2c).

FIGURE 2.

Engineering and physicochemical characterization of a bioactive conjugate‐derived nanotherapy, TPT NP, and its in vitro biological effects. (a,b) Typical TEM image (a) and size distribution (b) of TPT NP. (c) Dose‐dependent elimination of superoxide anion, DPPH/hydroxyl radicals, and H₂O₂ by TPT NP. (d) A Lineweaver–Burk plot used to analyze the effects of TPT NP on the kinetics of H₂O₂ catalysis and scavenging. The black dashed lines represent linearly fitted curves based on experimental data, which were used to quantify and compare the redox buffering capacities of the two groups. (e) The effect of freeze–drying cycle number on the hydrodynamic diameter of TPT NP. (f,g) TEM images of TPT NP after the first freeze–drying cycle (f) and after the final freeze–drying process (g). (h) ¹H NMR spectrum of hydrolyzed products of PBAP after incubation with H₂O₂. (i) Representative fluorescence images showing time‐dependent cellular uptake of Cy5‐TPT NP (400 µg/mL) in hUSMCs. (j) Fluorescence microscopic images illustrate dose‐dependent cellular uptake of Cy5‐TPT NP in hUSMCs after 2 h of incubation. (k) qPCR quantification of mRNA levels of TNF‐α, IL‐1β, and IL‐6 in hUSMCs after treatment with different formulations. Control, cells treated with medium alone; Model, cells stimulated with LPS; TPT NP groups, cells stimulated with LPS and treated with different doses of TPT NP; Atosiban, cells stimulated with LPS and treated with atosiban at 0.27 µg/mL. (l,m) Representative fluorescence images of DHE‐stained hUSMCs (l) and SOD enzyme activity in hUSMCs (m) after pre‐treatment with different doses of TPT NP or atosiban and stimulation with H₂O₂. (n) Typical flow cytometric curves of DHE‐stained hUSMCs after pre‐treatment with TPT NP (0.4 mg/mL) or PEG (0.32 mg/mL) and stimulation with H₂O₂. Data in (c,e,k,m) are mean ± s.d. (n = 6 independent samples). Statistical significance was assessed by one‐way ANOVA with the post hoc LSD test. *p < 0.05, **p < 0.01, ***p < 0.001.

Accumulating evidence indicates that PTB is closely associated with elevated ROS levels and impaired redox homeostasis [49]. We therefore evaluated the redox buffering capacity of TPT NP. 2′,7′‐dichlorodihydrofluorescein (DCFH) assay was conducted, in which the fluorescence intensity of the oxidized product 2′,7′‐dichlorofluorescein (DCF) serves as a quantitative indicator of ROS levels in the reaction system. First, we assessed the in vitro redox buffering capacity of TPT NP in human USMCs (hUSMCs) under H₂O₂ stimulation. The DCFH oxidation rate increased with rising H₂O₂ concentrations. However, in the presence of 400 µg/mL TPT NP, fluorescence intensity showed minimal variation across H₂O₂ concentrations ranging from 25 to 200 µm, indicating effective stabilization of intracellular ROS levels. TPT NP did not completely scavenge ROS. Above 200 µm H₂O₂, the redox buffering capacity of TPT NP became saturated, as evidenced by a sharp increase in fluorescence intensity. This occurred because the excessive oxidative load exceeded the regulatory capacity of TPT NP, thereby disrupting redox homeostasis (Figure S2a). We next quantified the redox buffering capacity (β), defined as the change in oxidant or reductant concentrations required for a 1 V shift in effective reduction potential (β = δC_ox/δE_eff, where δC_ox = oxidant concentration change and δE_eff = reduction potential change). Experimentally, β was derived from the relationship: 1/ΔVf = 1/ΔVf _max + (kcap/ΔVf _max) × 1/C_{o x} (where ΔVf _max = maximum DCFH oxidation rate and kcap = dimensionless concentration‐dependent constant). β was calculated as the slope (β = kcap/ΔVf _max). This analysis revealed that TPT NP exhibited a significantly higher buffering capacity (∼540.6 units) than intrinsic cellular components (∼169.4 units), corresponding to a 3.2‐fold enhancement (Figure 2d). These results indicate that TPT NP possesses excellent redox buffering capacity, supporting their potential application for the prevention and/or treatment of PTB.

To further assess the practical applicability of TPT NP, we examined their stability and recyclability. The hydrodynamic diameter and ζ‐potential of TPT NP exhibited negligible variation over five freeze–drying/redispersion cycles, indicating good colloidal stability and structural integrity (Figure 2e; Figure S2b). TEM imaging further confirmed that particle morphology and size remained unchanged after repeated processing (Figure 2f,g). These findings demonstrate that TPT NP can be recovered and redispersed multiple times without significant aggregation or structural disruption, supporting their excellent recyclability. Meanwhile, considering the well‐documented anti‐inflammatory activity of HMP, the hydrolytic products of PBAP and TPT NP were characterized using ¹H NMR spectroscopy. It was found that PBAP undergoes ROS‐mediated conversion to HMP, which underlies the antioxidant and anti‐inflammatory effects of TPT NP as well. Specifically, new characteristic proton signals emerged at ∼6.8 ppm (aromatic protons) and ∼4.5 ppm (─CH₂─ group) (Figure 2h), consistent with chemical shifts of HMP. Concurrently, the original signals of PBAP exhibited distinct changes. As for TPT, proton signals corresponding to HMP were also detected in the presence of ROS, confirming HMP generation (Figure S2c). These results provide direct experimental evidence for the ROS‐responsive release of HMP from TPT NP.

2.2. Antioxidative and Anti‐Inflammatory Effects of TPT NP

Subsequently, we evaluated the biological activities of TPT NP in vitro. It has been well‐established that abnormal contraction of USMCs plays a key role in the occurrence of PTB [50, 51, 52]. Consequently, we investigated the effects of TPT NP on hUSMCs. To facilitate fluorescent tracking, a cyanine5 (Cy5)‐labeled conjugate, that is, Cy5‐TCP, was synthesized by covalently linking both Cy5 and PEG to TCT. Cy5‐labeled TPT NP (Cy5‐TPT NP) was then obtained by co‐assembling Cy5‐TCP and TPT at a 1:9 molar ratio. The resulting Cy5‐TPT NP exhibited a hydrodynamic diameter of 98 nm and ζ‐potential −3.4 ± 1.3 mV (Figure S3). Cellular uptake of Cy5‐TPT NP was examined by in vitro culture of hUSMCs. Confocal laser scanning microscopy (CLSM) observation showed that Cy5‐TPT NP could be effectively internalized in hUSMCs, demonstrating clear dose‐ and time‐dependent effects (Figure 2i,j), which was confirmed by flow cytometry analysis (Figure S4).

Excessive extracellular ROS production and massive pro‐inflammatory factor secretion are well‐recognized pathological features of PTB, which further trigger cell death and tissue damage [53]. Therefore, to evaluate anti‐inflammatory and anti‐oxidative effects of TPT NP, we detected the expression of inflammatory cytokines, including tumor necrosis factor (TNF)‐α, interleukin (IL)‐1β, and IL‐6, in lipopolysaccharide (LPS)‐stimulated hUSMCs, a well‐established cell model to assess inflammatory response in PTB [54]. qPCR quantification revealed that LPS simulation significantly increased mRNA levels of TNF‐α, IL‐1β, and IL‐6 (Figure 2k). TPT NP treatment dose‐dependently downregulated the expression of these cytokines. In particular, 400 µg/mL TPT NP restored mRNA levels of these pro‐inflammatory cytokines to those comparable with the normal control group. Similarly, in an H₂O₂‐induced oxidative stress model [36], TPT NP at all three tested doses significantly reduced intracellular ROS generation and enhanced SOD enzymatic activity in hUSMCs, as confirmed by both CLSM observation and flow cytometric quantification (Figure 2l,m; Figure S5a–c). Notably, TPT NP demonstrated significantly superior efficacy compared to atosiban, a commonly used clinical tocolytic for PTB prevention [55]. It has been reported that PEG itself possesses certain antioxidant properties [56]. To assess the intrinsic antioxidant contribution of PEG, an equimolar control experiment was conducted comparing free PEG with TPT NP containing an equivalent molar amount of PEG. The results showed that PEG alone induced only a modest, statistically insignificant decrease in intracellular ROS levels, while TPT NP exhibited significantly stronger ROS‐scavenging activity (Figure 2n; Figure S5d–f). These findings suggest that PEG itself contributes minimally to the overall antioxidant effect, and the potent antioxidant activity of TPT NP stems from the synergistic interplay of Tempol, PBAP, and PEG. Taken together, these results substantiate that TPT NP exhibits potent anti‐inflammatory and antioxidant activities in stimulated hUSMCs, highlighting its potential as a novel therapy for PTB intervention by simultaneously regulating oxidative stress and inflammation.

2.3. Amelioration of Abnormal Contraction and Apoptosis of hUSMCs by TPT NP

Uterine smooth muscle exhibits abnormal contractions during PTB, a pathological process closely linked to massive Ca²⁺ influx through sarcolemmal calcium channels [57, 58]. Using the calcium‐sensitive dye Fura‐4 AM, we observed that LPS induction significantly increased intracellular Ca²⁺ fluorescence intensity in hUSMCs, compared to the control cells. Treatment with TPT NP at varying concentrations progressively reduced this signal (Figure 3a; Figure S6a), confirming its efficacy in counteracting calcium dysregulation.

FIGURE 3.

Multiple bioactivities of TPT NP in hUSMCs pathologically relevant to PTB. (a) Representative fluorescence images reveal intracellular Ca²⁺ levels in hUSMCs after 24 h of incubation with different concentrations of TPT NP or atosiban. (b,c) Representative fluorescence images (b) and western blot analysis (c) indicate Cx43 protein levels in hUSMCs after different treatments. (d,e) Representative flow cytometric profiles (d) and quantitative analysis (e) of apoptotic hUSMCs. (f) Representative fluorescence images show relative levels of cleaved caspase‐3 in hUSMCs subjected to different treatments. Control, cells treated with medium alone; Model, cells stimulated with H₂O₂; TPT NP groups, cells stimulated with H₂O₂ and treated with different doses of TPT NP; Atosiban, cells stimulated with H₂O₂ and treated with atosiban. (g) Representative gel contraction photographs of hUSMCs. Images show the control group, model (LPS) group, LPS plus different concentrations of TPT NP group, and LPS plus atosiban group at 0, 24, and 48 h post‐treatment. The yellow‐dotted lines highlight the extent of gel contraction. Data in (e) are mean ± s.d. (n = 6 independent samples). Statistical significance was assessed by one‐way ANOVA with the post hoc LSD test. *p < 0.05, ***p < 0.001.

Connexin 43 (Cx43), an established biomarker of cellular contractility, was significantly upregulated in LPS‐treated hUSMCs, indicating enhanced contractile activity [59, 60]. As illustrated in Figure 3b,c; Figure S6b, TPT NP treatment effectively reversed this elevated Cx43 expression, with the efficacy of TPT NP at 400 µg/mL significantly surpassing that of atosiban, a known oxytocin receptor antagonist [61]. Previous studies have shown that sustained calcium overload is known to disrupt intracellular homeostasis by activating calcium‐dependent apoptotic enzymes (e.g., caspases), ultimately triggering apoptosis [62, 63, 64, 65]. Aberrant Cx43 overexpression not only drives contraction but also sensitizes cells to apoptotic stimuli via downstream signaling pathways [66, 67]. We therefore assessed apoptosis of H₂O₂‐stimulated hUSMCs using flow cytometry and immunofluorescence following different treatments. It was found that TPT NP treatment significantly reduced apoptosis compared to the H₂O₂‐induced model group (Figure 3d–f).

More direct functional evidence was provided by collagen gel contraction assays. LPS induction caused notable contraction in the model group, which was markedly suppressed by TPT NP treatment. Compared to the control, the gel area was significantly reduced in the LPS group at both 24 and 48 h. In contrast, TPT NP dose‐dependently restored the gel area, with 400 µg/mL TPT NP outperforming atosiban at corresponding time points (Figure 3g; Figure S6c). Collectively, these findings demonstrate that TPT NP can effectively attenuate abnormal contraction and apoptosis in hUSMCs, primarily by lowering intracellular calcium levels and downregulating Cx43 expression.

2.4. Tissue Distribution Profiles of TPT NP in Mice With LPS‐Induced PTB

Before in vivo therapeutic studies, we first examined in vivo biodistribution profiles of TPT NP in mice with LPS‐induced PTB (Figure 4a). In vivo fluorescence imaging revealed time‐dependent distribution of Cy5‐TPT NP (at 10 mg/kg) in the liver and uterus within 24 h after intravenous (i.v.) injection (Figure 4b,c), indicating its ability to accumulate at uterine inflammation sites. These results provide direct evidence for uterine‐selective accumulation of TPT NP from systemic circulation, establishing a pharmacokinetic rationale for its therapeutic efficacy. Furthermore, ex vivo fluorescence imaging of harvested organs confirmed predominant uterine localization, with fluorescence intensity gradually increasing over 0–24 h (Figure 4d,e). It is worth noting that fluorescence in the placenta was primarily localized in the maternal decidual layer, without penetration through the placental barrier (Figure 4d–g). Importantly, no fluorescence was detected in fetal tissues, while maternal accumulation occurred mainly in the liver, lung, and kidney (Figure 4h). Together, these results indicate that TPT NP can accumulate in the uterus and placental decidua of PTB mice, without crossing the placental barrier, thereby minimizing fetal exposure.

FIGURE 4.

Tissue distribution profiles of TPT NP in pregnant mice with LPS‐induced PTB after i.v. injection. (a) Treatment protocols for in vivo distribution studies of Cy5‐TPT NP in PTB mice. (b,c) Representative in vivo fluorescence images show the accumulation of Cy5‐TPT NP in PTB mice (E16.5) at different time points after i.v. injection (b) and quantified fluorescence intensities (c). (d–f) Ex vivo fluorescence images show the accumulation of Cy5‐TPT NP in the uterus and placenta of PTB mice (E16.5) at predetermined time points after i.v. injection (d) and quantified fluorescence intensities in uterine (e) and placental (f) tissues. (g) Representative fluorescence whole‐slide images of placental sections after i.v. injection of saline or Cy5‐TPT NP in PTB mice. (h) The accumulation of Cy5‐TPT NP in the maternal major organs (heart, liver, spleen, lung, and kidney) and fetuses at 24 h after i.v. injection in PTB mice. Data in (c,e,f) are mean ± s.d. (n = 6 independent mice).

2.5. TPT NP Delays LPS‐Induced PTB and Attenuates Adverse Pregnancy Outcomes

Following confirmation of the uterine‐targeting capability of TPT NP, we evaluated its therapeutic effects in mice with LPS‐induced PTB. At 0.5 h after LPS challenge, mice in TPT NP groups were treated with TPT NP at 2, 5, and 10 mg/kg via i.v. administration, while mice treated with atosiban (0.17 mg/kg, a clinically relevant dose) served as the positive control (Figure 5a). All animals in the normal control group delivered at full term (72 h after saline injection), whereas every mouse in the LPS‐induced model group delivered prematurely within 36 h (Figure 5b). TPT NP treatment delayed the onset of PTB in a dose‐dependent manner. By contrast, all atosiban‐treated mice were delivered within 48 h. Critically, TPT NP at 10 mg/kg significantly outperformed atosiban in prolonging gestational duration. We also evaluated postnatal pup survival across treatment groups, observing a markedly low survival rate in the model group, which was improved by TPT NP in a dose‐response pattern (Figure 5c). The 10 mg/kg TPT NP group yielded significantly higher offspring survival rates than atosiban. These results collectively demonstrate the dose‐dependent efficacy of TPT NP in delaying PTB and enhancing neonatal survival.

FIGURE 5.

Inhibition of LPS‐induced PTB in pregnant mice by TPT NP. (a) Schematic diagram illustrating the treatment protocol for therapeutic studies. (b) Changes in pregnancy rates over time in normal and PTB pregnant mice treated with different formulations. (c) Survival rate of mouse offspring after delivery. (d–f) Representative digital photographs show fetal morphology (d), with quantitative analyses of fetal weight (e) and crown‐rump length (CRL) (f) following different treatments. Box plots illustrate data distributions (n = 6 pregnant mice), with the central lines representing the mean, box boundaries indicating standard deviation, and whiskers showing the minimum and maximum values. (g) Micrographs of H&E‐stained histological sections of fetuses at birth. (h) Alizarin red and Alcian blue staining of the entire skeleton of fetuses from different groups. (i) Whole‐slide images (top) and high‐magnification images (lower) of H&E‐stained placental sections. Black dashed lines indicate the spongy trophoblast layer, while red and purple dashed lines denote the labyrinth layer and the maximum invasion distance, respectively. (j) Changes in pregnancy rates over time in normal and PTB pregnant mice treated with TPT NP at 10 mg/kg, atosiban at 0.17 mg/kg, or indomethacin at 1 mg/kg. Data in (c) are mean ± s.d. (n = 6 pregnant mice). Statistical significance was assessed by one‐way ANOVA with the post hoc LSD test. **p < 0.01, ***p < 0.001.

To assess whether TPT NP can ameliorate PTB‐associated adverse pregnancy outcomes, pregnant mice at E16.5 were challenged with LPS, followed by i.v. administration of TPT NP at various doses (2, 5, and 10 mg/kg) or 0.17 mg/kg atosiban 30 min later. All mice were euthanized at delivery, and fetuses and placentas were subsequently collected. It was found that the model group exhibited significantly reduced fetal weight compared to the control group (Figure 5d,e). Both TPT NP and atosiban treatments effectively restored fetal weight, with the 10 mg/kg TPT NP dose showing superior efficacy to atosiban. Similarly, crown‐rump length (CRL), a key metric of fetal growth, was significantly shortened in the model group (Figure 5f). TPT NP treatment effectively increased CRL, again showing superior efficacy for the 10 mg/kg dose compared to atosiban. These findings on fetal weight and CRL were corroborated by histological analysis of H&E‐stained fetal sections (Figure 5g). Additionally, fetal skeletal development was evaluated. Compared to the control group, fetuses from the model group exhibited a higher proportion of cartilage (Alcian blue staining) and a marked reduction in bone tissue (Alizarin red staining), along with occipital hypoplasia, indicating a substantial delay in overall fetal development. These skeletal abnormalities were markedly ameliorated by TPT NP treatment, with the 10 mg/kg dose showing the most pronounced effect (Figure 5h).

The placenta, the central organ for maternal–fetal exchange, is vital for supplying oxygen and nutrients while removing metabolic wastes, thereby supporting normal fetal growth and development [68]. Intrauterine inflammation, however, can disrupt these critical functions. It not only impairs placental morphogenesis and functional maturation but may also disrupt the placental–embryonic blood supply balance, ultimately compromising fetal nutrient acquisition and the intrauterine environment [69]. Compared with the control group, mice in the model group exhibited significantly reduced placental weights. In contrast, both TPT NP and atosiban treatment effectively restored placental weights (Figure S7a,b). Additionally, the model group exhibited marked thickening of the spongiotrophoblast layer and increased trophoblast invasion distance, indicating that trophoblast cells excessively invaded the decidua. Meanwhile, the labyrinth layer became thinner. This structural abnormality is associated with insufficient vascular branching in the placental labyrinth layer, thereby impairing embryonic growth. Critically, these placental abnormalities were reversed following TPT NP treatment (Figure 5i; Figure S7c,d).

Indomethacin, a nonsteroidal anti‐inflammatory drug, can effectively inhibit uterine contraction in the short term by suppressing prostaglandin synthesis [70, 71]. It is mainly used to suppress preterm labor contractions before 32 weeks of gestation. However, its clinical application is limited by risks such as premature closure of the fetal ductus arteriosus [72]. We further examined pregnancy rates throughout gestation and used indomethacin (1 mg/kg) as a positive control for parallel comparison (Figure 5j). Indomethacin showed minimal therapeutic efficacy in this model, with a pregnancy rate profile comparable to that of the model group, indicating that it cannot delay delivery once uterine contractions have been initiated. In contrast, atosiban effectively prolonged gestation and exerted obvious anti‐PTB effects. Notably, 10 mg/kg TPT NP outperformed both atosiban and indomethacin in delaying PTB progression and sustaining pregnancy. These findings demonstrate that TPT NP possesses superior therapeutic potential in a clinically relevant PTB model characterized by inflammation and initiated uterine contractions, providing a promising strategy for PTB management.

Collectively, our findings demonstrate that the multi‐bioactive nanotherapy TPT NP can effectively prevent LPS‐induced PTB, improve offspring survival, promote fetal growth, and ameliorate placental pathological abnormalities associated with oxidative stress and inflammation. Notably, the therapeutic efficacy of TPT NP at 10 mg/kg was markedly superior to that of atosiban at a clinically relevant dose.

2.6. TPT NP Preserves Maternal–Fetal Interface Homeostasis via Potent Anti‐Inflammatory Effects in PTB Mice

The maternal–fetal interface, comprising the uterus, placenta, and fetal membranes, is essential for maintaining pregnancy homeostasis. Disruption of the inflammatory and redox balance at this interface can lead to adverse pregnancy outcomes. LPS, a key component of the gram‐negative bacterial cell wall and a potent pro‐inflammatory agent, has been extensively shown to disrupt the maternal–fetal interface [73, 74]. Building on the above promising therapeutic effects of TPT NP in mice with LPS‐induced PTB, we further explored the underlying mechanisms, focusing on its regulation of inflammatory imbalance at the maternal–fetal interface. To evaluate systemic inflammation, a LPS‐induced PTB model was established in E16.5 mice. At 8 h after i.v. administration of TPT NP or atosiban, peripheral blood was collected for white blood cell (WBC) counting. The LPS model group exhibited markedly elevated WBC counts, validating the successful establishment of systemic inflammation. TPT NP treatment significantly reduced WBC levels in a concentration‐dependent manner, demonstrating potent anti‐inflammatory activity. In contrast, atosiban exerted no significant effect on WBC counts compared with the LPS group, confirming that it protects against PTB only via inhibiting uterine contractions and lacks anti‐inflammatory activity (Figure S8). Furthermore, immunohistochemistry and qPCR analyses showed that TPT NP potently suppressed LPS‐induced upregulation of TNF‐α, IL‐1β, and IL‐6 in uterine, placental, and fetal membrane tissues (Figure 6a–d; Figure S9). Conversely, atosiban, which acts exclusively through inhibiting uterine contractions, displayed no anti‐inflammatory activity. These results confirm that TPT NP exhibits potent anti‐inflammatory effects at the maternal–fetal interface.

FIGURE 6.

Anti‐inflammatory and anti‐oxidative effects of TPT NP in LPS‐induced PTB mice. (a–d) Representative immunohistochemical images indicating relative levels of IL‐6 in uterine, placental, and fetal membrane (FM) tissues (a), along with quantitative analysis of mRNA levels of TNF‐α (b), IL‐1β (c), and IL‐6 (d) in uterine tissues. (e) Representative fluorescent images of uterine, placental, and fetal membrane tissues stained with DHE. (f) Immunofluorescence images illustrate relative levels of MPO. For these studies, LPS‐induced PTB in pregnant mice was established at E16.5. After LPS stimulation for 30 min, both the sham surgery (control) and model groups received saline by i.v. injection, while the TPT NP groups were treated with TPT NP at 2, 5, or 10 mg/kg. The atosiban group received i.v. injection of atosiban at 0.17 mg/kg. Mice in all groups were euthanized at 8 h after different treatments, and relevant samples were collected for evaluations. Data in (b–d) are mean ± s.d. (n = 6 independent mice). Statistical significance was assessed by one‐way ANOVA with Tamhane's T2 test (b,c) or post hoc LSD test (d). *p < 0.05, **p < 0.01, ***p < 0.001.

Inflammatory responses and oxidative stress establish a self‐amplifying vicious cycle. LPS‐induced PTB triggers inflammatory factor release and activates immune cells like macrophages and neutrophils, thereby stimulating excessive ROS production that causes uterine oxidative damage [75, 76]. Accordingly, we evaluated oxidative stress‐related parameters, including ROS levels (detected by a fluorescence probe dihydroethidium (DHE)) and neutrophil infiltration (assessed by MPO staining). The results indicated that TPT NP significantly attenuated LPS‐induced ROS accumulation and excessive neutrophil infiltration in uterine, placental, and fetal membrane tissues (Figure 6e,f; Figure S10). Furthermore, LPS challenge disrupted redox balance in gestational tissues, resulting in substantial H₂O₂ accumulation and elevated malondialdehyde (MDA), a lipid peroxidation marker reflecting the extent of oxidative membrane damage [77, 78]. Meanwhile, excessive ROS leads to the depletion of key antioxidant enzymes, including SOD, catalase (CAT), and glutathione peroxidase (GPx), resulting in a pronounced reduction in their enzymatic activities [79, 80]. More importantly, the total equivalent antioxidant capacity (TEAC) of tissues also decreases significantly, further aggravating the oxidative stress imbalance [81]. Notably, treatment with TPT NP effectively restored these oxidative stress markers, with efficacy superior to that of atosiban (Figures S11,S12). These results confirm that TPT NP effectively restores redox homeostasis in LPS‐induced PTB, primarily by reducing oxidative damage products and enhancing endogenous antioxidant defenses. This dual protective mechanism breaks the vicious cycle between inflammation and oxidative stress, thereby preserving maternal–fetal interface homeostasis.

2.7. TPT NP Improves Blood Flow Abnormality and Uterine Contraction in PTB Mice

Intrauterine inflammation and oxidative stress are central pathological events in PTB, largely mediated by placental perfusion dysfunction and hemodynamic imbalance. This dysfunction acts both as a direct result of inflammatory injury and as a crucial intermediary in conveying maternal inflammatory signals to the fetus, ultimately contributing to PTB [82, 83, 84]. To evaluate the effect of TPT NP on placental perfusion deficits, we induced a PTB model by injecting 20 µg LPS between the first and second amniotic sacs in the lower portion of the right uterine horn of each healthy pregnant mouse at embryonic day 16.5 (E16.5), followed by i.v. administration of TPT NP (2, 5, or 10 mg/kg) at 30 min post‐LPS challenge. A positive control group received atosiban at 0.17 mg/kg, while a sham‐operated control group was treated with saline alone to minimize procedural and solvent interferences.

Ultrasound color Doppler was used to non‐invasively monitor hemodynamic parameters, including the resistance index (RI), pulse index (PI), and systolic/diastolic ratio (S/D), which reflect vascular resistance, pulsatility, and perfusion efficiency, respectively [85, 86]. Compared with the control and TPT NP groups, mice in the model group showed significantly elevated uterine and umbilical artery RI, PI, and S/D values, indicating increased vascular resistance and impaired vasodilation due to inflammation (Figure 7a–d). This led to reduced placental blood flow and compromised fetal nutrient–oxygen supply. Critically, TPT NP treatment dose‐dependently reversed these abnormalities. At 10 mg/kg TPT NP, the mentioned parameters were restored to near‐normal levels, confirming the mitigation of ROS‐ and inflammation‐mediated vascular dysfunction. These in vivo results align with prior in vitro evidence that TPT NP possesses potent anti‐inflammatory and anti‐oxidative efficacy.

FIGURE 7.

Mechanisms underlying therapeutic effects of TPT NP in treating LPS‐induced PTB in pregnant mice. (a–d) Doppler blood flow waveforms of the uterine artery (a) and fetal umbilical artery (c), along with quantitative analysis of the resistance index (RI), pulse index (PI), and systolic/diastolic flow velocity ratio (S/D) for the uterine artery (b) and fetal umbilical artery (d). (e) Representative immunofluorescence images showing relative expression levels of Cx43 in cryosections of uterine tissues after different treatments. The dose of atosiban was 0.17 mg/kg. (f,g) Typical Western blot bands (f) and quantitative analysis (g) indicate Cx43 protein levels in uterine tissues following different treatments. (h) Quantified values of the mean muscle tension, maximum contraction force, and area under the curve (AUC) for uterine muscle contraction curves after different treatments. In these studies, the LPS‐induced PTB model in pregnant mice was established at E16.5. After LPS stimulation for 30 min, both the sham surgery (control) and model groups received saline by i.v. injection, while the TPT NP groups were treated with TPT NP at 2, 5, or 10 mg/kg. The atosiban group received i.v. injection of atosiban at 0.17 mg/kg. Mice in all groups were euthanized at 8 h after different treatments, and relevant samples were collected for evaluations. Data in (b,d,g,h) are mean ± s.d. (n = 6 independent mice). Statistical significance was assessed by one‐way ANOVA with the post hoc LSD test (b,d,g) or Tamhane's T2 test (h). *p < 0.05, **p < 0.01, ***p < 0.001.

Following an 8 h intervention with various doses of TPT NP or atosiban in PTB mice, uterine tissues were collected for immunofluorescence and Western blot analyses. The results showed that TPT NP significantly downregulated the expression of Cx43, a key regulatory protein of uterine contractions (Figure 7e–g). The abnormal expression of Cx43 is closely associated with excessive uterine smooth muscle contraction. Consequently, the regulatory effect of TPT NP on this protein further elucidates its potential molecular mechanism for ameliorating pathological conditions during pregnancy. Real‐time uterine contraction monitoring using high‐precision tension sensors revealed that TPT NP treatment significantly reduced average and peak contractile forces, along with cumulative contractile activity (quantified as AUC) (Figure 7h; Figure S13). Similarly, atosiban, a classic oxytocin receptor antagonist, also markedly suppressed uterine contractility.

2.8. Safety Studies

Finally, safety profiles of TPT NP were evaluated. Initially, we assessed the cytotoxicity of the TCT scaffold toward hUSMCs, and the results showed that TCT exerted significant cytotoxicity at concentrations higher than 64 µg/mL. In contrast, TPT, prepared by substituting the three chlorine atoms in TCT, exhibited no significant cytotoxic effects on hUSMCs even at concentrations up to 1 mg/mL (Figure S14a,b). This observation is consistent with the fact that the cytotoxicity of TCT primarily attributed to its C─Cl bonds. Specifically, due to the strong electron‐withdrawing effect of chlorine atoms, the carbon atom in the C─Cl bond exhibits significant electrophilicity, making it susceptible to attack by electron‐rich nucleophilic groups (e.g., sulfhydryl and amino groups) in biomolecules, thereby triggering nucleophilic substitution reactions [87, 88]. These reactions can induce alkylation modifications in proteins, leading to their inactivation and disruption of spatial conformation. Simultaneously, the C─Cl bonds cause DNA base alkylation and strand breaks, ultimately resulting in cellular structural damage and genetic material destruction.

Also, in vitro tests suggested that TPT NP exhibited negligible hemolysis at different concentrations examined (Figure S14c,d). To evaluate the in vivo safety of TPT NP, pregnant mice were administered high doses (50 and 100 mg/kg, i.e., 5 and 10‐fold the therapeutic dose, respectively) daily from E10.5 for 8 days. All animals remained healthy, showing normal behaviors and gestational weight gains comparable to the saline control (Figure S15a). No significant differences were observed in the organ‐to‐body weight ratios of maternal major organs, fetuses, or placentas (Figure S15b,c). Gross examination following H&E staining revealed normal fetal and placental morphology with negligible malformations (Figure 8a,b). Histopathological analysis showed no tissue damage or inflammatory infiltration in maternal or fetal organs (Figure 8c,d). Furthermore, hematological and biochemical parameters (relevant to liver and kidney functions) in maternal blood samples were within normal ranges (Figure S16), collectively demonstrating a favorable safety profile of TPT NP in pregnant mice.

FIGURE 8.

Safety evaluations of TPT NP in normal pregnant mice. (a,b) H&E‐stained histological sections of fetuses (a) and placentas (b) from pregnant mice after different treatments. (c,d) Microscopic images illustrate H&E‐stained sections of major organs from maternal mice (c) and fetuses (d). In these studies, pregnant mice at E10.5 received daily i.v. injection of TPT NP (at doses of 50 and 100 mg/kg) for 8 days. For the control group, pregnant mice were treated with saline. At E18.5 after different treatments, all treated pregnant mice were euthanized. The fetuses, placentas, and typical major organs, including the heart, liver, spleen, lung, and kidney were collected from either maternal mice or fetuses for analyses.

Considering the crucial role of uterine immune homeostasis in maintaining pregnancy and ensuring i.v. treatment safety, we assessed whether i.v. administration of TPT NP triggers discernible immune responses. Evaluations were performed in female mice at both acute (24 h) and chronic (30 days) time points post‐administration. Quantitative analysis of plasma samples showed that the levels of the pro‐inflammatory mediators TNF‐α and IL‐6 in the TPT NP‐treated group did not differ significantly from those in the control group at either time point, and no marked inflammatory responses were observed (Figure S17). Histopathological examination indicated that neutrophil and macrophage infiltration in uterine tissues was comparable between the TPT NP and control groups at 24 h and 30 days (Figure 9a,b). Furthermore, the abundance of CD3⁺ lymphocytes in uterine and splenic tissues showed no significant differences between groups (Figure 9c,d). Consistent with these results, TPT NP administration did not alter the organ indices of the uterus or spleen (Figure S18), ruling out treatment‐related tissue edema. Together, these preliminary findings substantiate that TPT NP does not induce acute or chronic immune activation in mice.

FIGURE 9.

Evaluation of potential inflammatory and immune responses after TPT NP treatment in pregnant mice. (a) Representative immunofluorescence images showing relative expression levels of MPO⁺ neutrophils in cryosections of uterine tissues. (b,c) Immunohistochemical images show relative counts of F4/80⁺ macrophages (b) and CD3⁺ lymphocytes (c) in uterine sections. (d) Immunohistochemical staining of CD3⁺ lymphocytes in spleen sections. In these experiments, normal pregnant mice received i.v. injection of either saline (control) or TPT NP (at 50 or 100 mg/kg). Uterine and spleen tissues were collected from the dams at 24 h or day 30 post‐intervention for different analyses.

3. Discussion

PTB, a prevalent clinical obstetric complication, is pathologically mediated by excessive oxidative stress activation and inflammatory imbalance during pregnancy [89, 90]. Current clinical interventions exhibit substantial limitations, primarily relying on symptomatic management. These approaches demonstrate restricted efficacy in delaying PTB progression and improving adverse neonatal outcomes, with no definitive curative therapies established to date [55, 91]. Targeted nanotherapies have emerged as a prominent therapeutic research focus for the treatment of gynecological diseases and pregnancy‐associated diseases [92]. Consequently, developing nanotherapeutics capable of simultaneously regulating oxidative stress and inflammatory responses represents a promising strategy for optimizing PTB management.

In response to this clinical imperative and research direction, we propose a novel conjugate‐based nanotherapeutic strategy employing multi‐bioactive nanoparticles to target PTB pathology. We rationally designed an amphiphilic conjugate based on the TCT molecular skeleton, covalently integrating hydrophilic/biocompatible PEG with pharmacologically active functional modules: a SOD mimetic and a moiety capable of scavenging H₂O₂ and generating anti‐inflammatory compounds. This synthesis successfully conferred amphiphilicity, ROS‐responsive/scavenging capability, and anti‐inflammatory functionality, enabling the conjugate to self‐assemble into micelle‐like nanoparticles (TPT NP). The resulting TPT NP nanotherapy, with a uniform spherical morphology and an average diameter of 100 nm, exhibits potent antioxidant and anti‐inflammatory effects. In vitro assessments confirmed its dose‐dependent capacity to eliminate superoxide anions, DPPH, and hydroxyl radicals, and H₂O₂. Moreover, TPT NP showed outstanding recyclability and robust redox buffering capability, thereby laying a solid foundation for the effective treatment of PTB.

To further validate the effects of TPT NP on pathological cells associated with PTB and elucidate the underlying mechanisms, we performed comprehensive studies in hUSMCs. At the cellular level, the mechanism of TPT NP action primarily stems from its highly efficient intracellular delivery capacity. TPT NP demonstrated highly efficient, dose‐ and time‐dependent cellular uptake, establishing a foundation for targeted intracellular drug delivery. Crucially, TPT NP simultaneously inhibited oxidative stress and inflammatory cascades, addressing the core pathological drivers of PTB progression. Moreover, TPT NP effectively reduced LPS‐induced calcium ion influx in hUSMCs, thereby directly inhibiting cellular contraction. Concurrently, TPT NP significantly downregulated aberrantly high expression of the contraction marker Cx43, thus disrupting contraction signal transmission. This dual mechanism synergistically suppresses pathological uterine contractility by modulating the Ca²⁺‐Cx43 pathway axis. Furthermore, TPT NP significantly attenuated apoptosis in hUSMCs. These results collectively substantiate that TPT NP not only inhibits pathological uterine contractions but also promotes smooth muscle cell survival, thereby preserving normal uterine physiological function.

Given these promising cellular‐level findings, we next evaluated the in vivo targeting efficiency of TPT NP, as targeted delivery is pivotal for enhancing the efficacy of PTB therapies while minimizing systemic toxicity [36]. Using in vivo fluorescence imaging, we demonstrated that intravenously administered Cy5‐TPT NP preferentially accumulated at high concentrations in uterine and placental tissues, with no detectable uptake in fetal compartments. In terms of therapeutic efficacy, TPT NP demonstrated a significant dose‐dependent advantage. Compared with the model group, 10 mg/kg TPT NP significantly delayed LPS‐induced preterm labor in mice, improved offspring survival rate, and fetal/placental weights. Notably, TPT NP outperformed the clinical drugs atosiban and indomethacin in these metrics. Moreover, TPT NP effectively inhibited pathological trophoblast over‐invasion, restored the placental labyrinth structure, and promoted fetal skeletal development, advantages not afforded by contraction‐targeting drugs like atosiban. Mechanistically, TPT NP can simultaneously ameliorate the local oxidative‐inflammatory microenvironment in the placenta, thereby ensuring adequate nutrition and oxygen supply for fetal development. At the hemodynamic level, TPT NP reverses LPS‐induced abnormalities in uterine and fetal umbilical arterial blood flow, restoring placental–fetal perfusion and establishing a direct pathophysiological link between placental function and fetal development. Furthermore, both in vitro and in vivo findings substantiate that TPT NP may attenuate LPS‐induced uterine contractions. Crucially, i.v. administration of TPT NP at a 10‐fold therapeutic dose demonstrated excellent safety profiles in both maternal mice and offspring. Together, these findings establish a new paradigm for the safe, targeted treatment of PTB through advanced nanotherapy.

While TPT NP demonstrates promising therapeutic potential, this study has limitations. Notably, the LPS‐induced PTB model replicates only 25%–40% of clinical PTB cases, specifically those driven by infectious etiologies like chorioamnionitis. This model excludes other major contributors, such as cervical insufficiency, multiple gestation, and diverse non‐infectious pathways [93, 94]. Consequently, the broad applicability of TPT NP requires validation in a wider spectrum of established PTB models in future studies. Furthermore, optimizing delivery strategies or exploring combination therapies with existing agents could significantly extend the therapeutic potential of TPT NP to address multifactorial PTB.

Collectively, TPT NP demonstrates excellent recycling stability and redox‐buffering capacity. It specifically targets and accumulates in uterine and placental tissues, where it scavenges ROS and suppresses inflammatory responses. Through this core mechanism, TPT NP regulates the Ca²⁺‐Cx43 signaling axis to inhibit pathological uterine contractions, enhance placental function and perfusion, and maintain smooth muscle cell viability, thereby achieving safe and effective intervention against PTB. This work provides an innovative targeted nanotherapeutic strategy for the clinical management of PTB.

4. Conclusions

In this study, considering that existing clinical treatments for preterm labor only alleviate symptoms by suppressing uterine contractions without addressing the underlying causes of oxidative stress and inflammation, we designed and fabricated a multifunctional nanomedicine, TPT NP, which integrates antioxidant, anti‐inflammatory, and uterine contraction‐inhibiting activities. TPT NP is self‐assembled from the amphiphilic multifunctional conjugate TPT, exhibiting excellent broad‐spectrum ROS‐scavenging capability, redox‐buffering properties, and colloidal stability. In vitro studies demonstrated that TPT NP can be efficiently internalized by hUSMCs. By regulating the Ca²⁺‐Cx43 signaling pathway, it markedly suppresses abnormal cell contraction and apoptosis, while effectively blocking the cascade amplification of inflammation and oxidative stress. Its core therapeutic efficacy is significantly superior to that of atosiban, the first‐line clinical tocolytic agent. Furthermore, in vivo animal experiments confirmed that TPT NP specifically accumulates in inflamed uterine and placental tissues without crossing the placental barrier, thereby avoiding potential fetal development risks. It effectively delays LPS‐induced PTB in mice, significantly improves offspring survival rates, alleviates fetal growth restriction and placental structural abnormalities, and restores uteroplacental blood flow homeostasis. Importantly, safety evaluations further verified that TPT NP possesses favorable biocompatibility, with no adverse effects on the health of pregnant mice, fetal development, or immune homeostasis. In summary, this study overcomes the limitations of current PTB therapies that rely solely on symptomatic management. It establishes a novel nanotherapeutic strategy targeting the core pathophysiological mechanisms of PTB, providing a promising translational approach for the clinical prevention and treatment of inflammation‐related PTB. Additionally, this work offers a new conceptual framework for the management of other pregnancy complications driven by oxidative stress and inflammation.

5. Experimental Section

5.1. Materials

Cyanuric chloride (TCT) and 1,4‐dioxane were purchased from Macklin Biochemical Technology Co., Ltd. (Shanghai, China). Methoxy polyethylene glycol amine with a molecular weight of 2000 Da (mPEG‐NH₂) and cyanine5 amine (Cy5‐NH₂) were obtained from Xi'an Ruixi Biological Technology Co., Ltd. (Xi'an, China). 4‐Amino‐2,2,6,6‐tetramethylpiperidine‐1‐oxyl (Tempol) was purchased from Shanghai Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). (4‐(4,4,5,5‐Tetramethyl‐1,3,2‐dioxaborolan‐2‐yl) phenyl) methanol, that is, phenylboronic acid pinacol ester (PBAP) and lipopolysaccharide (LPS) were purchased from Sigma‐Aldrich (U.S.A.). Dichloromethane (CH₂Cl₂), methanol, and N,N‐diisopropylethylamine (DIPEA) were purchased from Beijing Bailingwei Technology Co., Ltd. (Beijing, China). 4’6‐Diamidino‐2‐phenylindole (DAPI) and dihydroethidium (DHE) were obtained from Beyotime Biotechnology (Nantong, China). Annexin‐V‐FITC cell apoptosis detection kit, hydrogen peroxide (H₂O₂), malondialdehyde (MDA), superoxide dismutase (SOD), glutathione peroxidase (GPx), catalase (CAT), total equivalent antioxidant capacity (TEAC) assay kit, and Fluo‐4 AM were purchased from Beyotime Biotechnology Co., Ltd. (China). GAPDH (60004‐1‐Ig), connexin 43 (26980‐1‐AP), HRP‐conjugated secondary antibody, FITC‐conjugated secondary antibody (SA00003‐2), and Cy3‐conjugated secondary antibody (SA00009‐1) were purchased from Proteintech Group, Inc (U.S.A.). Mouse myeloperoxidase (MPO, GB11224), F4/80 (GB11027) antibodies, and CD3 antibody (GB150004) were purchased from Wuhan Servicebio Technology Co., Ltd (Wuhan, China). Rat tail type I collagen (C8062) was obtained from Solarbio (China). Atosiban was purchased from Harbin Jixianglong Biotechnology Co., Ltd. (Harbin, China). Indomethacin was obtained from MedChemExpress Co., Ltd. (Shanghai, China). All other reagents were commercially available and used as received.

5.2. Synthesis and Characterization of a Bioactive Conjugate TPT

TPT was synthesized by simultaneously conjugating mPEG‐NH₂, Tempol, and PBAP onto TCT. Specifically, TCT (462 mg) was dissolved in anhydrous dichloromethane (20 mL) and cooled to 0°C. To this solution, 2 g mPEG‐NH₂ and 0.5 mL of DIPEA were added. The reaction mixture was stirred at 0°C for 24 h. After completion, the solvent was evaporated under reduced pressure. The residue was redissolved in 100 mL of dichloromethane and washed with water (3 × 10 mL) in a separatory funnel. The combined organic layers were dried over anhydrous MgSO₄, filtered, and concentrated. The crude product was purified by silica gel column chromatography (CH₂Cl₂/MeOH = 15:1) to afford PEG‐conjugated TCT (TCT‐PEG) as a white solid (1.5 g). Subsequently, a mixture of TCT‐PEG (1.5 g) and Tempol (120 mg) was dissolved in anhydrous 1,4‐dioxane (10 mL) under a nitrogen atmosphere. DIPEA (0.5 mL) was added, and the reaction was stirred at 40°C for 12 h. Then, 234 mg PBAP was added, and the mixture was heated at 80°C for an additional 12 h under nitrogen. Upon completion, the solvent was removed under reduced pressure. The residue was taken up in dichloromethane (100 mL) and washed with water (3 × 10 mL). The organic phase was dried over anhydrous MgSO₄, filtered, and concentrated. The final product, TPT, was obtained as a brown solid after purification by silica gel column chromatography (CH₂Cl₂/MeOH = 15:1) and drying in vacuo.

To synthesize a Cy5‐labeled conjugate, TCT‐PEG (33 mg) and Cy5‐NH₂ (10 mg) were dissolved in 5 mL of 1,4‐dioxane, followed by the addition of 0.2 mL of DIPEA. The mixture was stirred at 40°C for 12 h under a nitrogen atmosphere. After reaction, the organic solvent was removed by rotary evaporation. The residue was redissolved in 5 mL of deionized water, centrifuged, and the supernatant was lyophilized to afford Cy5‐conjugated TCT‐PEG (defined as Cy5‐TCP).

For materials characterization, ¹H NMR spectra were recorded on a nuclear magnetic resonance (NMR) spectrometer operating at 600 MHz (DD2, Agilent). Fourier transform infrared (FT‐IR) spectra were acquired on a PerkinElmer FT‐IR spectrometer (100S).

5.3. Anti‐Inflammatory Effects of TPT NP in hUSMCs

hUSMCs were seeded at 5 × 10⁵ cells/well in a 6‐well plate and cultured overnight. Cells were treated with fresh medium containing different concentrations of TPT NP (100, 200, and 400 µg/mL) or atosiban (0.27 µg/mL) for 24 h. Then, the medium was replaced with fresh medium containing lipopolysaccharide (LPS) at 1 µg/mL and incubated for 8 h. The control group was treated with PBS only. For quantitative analysis using qPCR, total RNA was extracted from hUSMCs utilizing TRIzol reagent (Thermo Fisher Scientific, U.S.A.) and quantified via NanoDrop (Thermo Fisher Scientific, U.S.A.). Total RNA was reverse transcribed into cDNA using the EvoScript Universal cDNA Master Reagent Kit (Roche, Germany). The primers for different pro‐inflammatory cytokines were as follows: TNF‐α (forward, CTCTTCTGCCTGCTGCACTTTG; reverse, ATGGGCTACAGGCTTGTCACTC), IL‐1β (forward, CCACAGACCTTCCAGGAGAATG; reverse, GTGCAGTTCAGTGATCGTACAGG), IL‐6 (forward, AACATGTGTGAAAGCAGCAAAGA; reverse, CTCTGGCTTGTTCCTCACTACTC), and GAPDH (forward, GGAGTCCACTGGCGTCTTCA; reverse, GTCATGAGTCCTTCCACGATACC). GAPDH was used as an endogenous reference.

5.4. The Effect of TPT NP on Intracellular Ca²⁺ in hUSMCs

hUSMCs were seeded at a density of 2 × 10⁵ cells/well into a 24‐well plate and cultured overnight. The cells were treated with fresh medium containing different concentrations of TPT NP (100, 200, and 400 µg/mL) or atosiban (0.27 µg/mL) for 24 h. Next, the medium was replaced with 1 µg/mL LPS and incubated for 8 h. After removing the culture medium, cells were washed with PBS, followed by the addition of 5 µmol/L Fura‐4 AM (a Ca²⁺ ion fluorescent probe) and incubation for 30 min. Then the fluorescence signals were observed by CLSM. In another parallel experiment, hUSMCs were inoculated into 96‐well plates overnight and treated in the same manner as described above. Subsequently, the absorbance values at 340 nm were measured using a microplate reader (Tecan) to quantitatively analyze intracellular Ca²⁺ levels.

5.5. The Effects of TPT NP on hUSMCs Contraction and Apoptosis

To evaluate the effect of TPT NP on the contraction of hUSMCs, cells were seeded at 2 × 10⁵ cells/well in a 24‐well plate and cultured overnight. They were then treated with fresh medium containing TPT NP at graded concentrations (100, 200, and 400 µg/mL) or atosiban (0.27 µg/mL) for 24 h, followed by replacement with medium containing 1 µg/mL LPS and incubation for an additional 8 h. Cells were fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.3% Triton X‐100, and blocked with 5% BSA for 30 min. Subsequently, cells were incubated with a primary antibody against Cx43 (1:400) at 4°C overnight, then with a Cy3‐labeled fluorescent secondary antibody at 37°C for 30 min. Fluorescence signals were then assessed by fluorescence microscopy. In addition, hUSMC samples of different groups were lysed with RIPA buffer (Servicebio, Wuhan, China). Then, the total protein concentration was measured using the BCA protein quantification kit (Servicebio, Wuhan, China). The expression levels of Cx43 in the hUSMCs were detected in each group by Western blot and quantitative analysis. In a separate experiment, similar treatment procedures were followed, with the exception that the incubation medium was replaced with medium containing 200 µmol/L H₂O₂, and the primary antibody was switched to cleaved caspase‐3 (1:200). The fluorescence intensity was observed by fluorescence microscopy.

To evaluate the effect of TPT NP on apoptosis of hUSMCs, cells were seeded at 2 × 10⁵ cells/well in a 24‐well plate and cultured overnight. Cells were then treated with fresh medium containing TPT NP at graded concentrations (100, 200, and 400 µg/mL) or atosiban (0.27 µg/mL) for 24 h, followed by replacement with medium containing 200 µmol/L H₂O₂ and incubation for an additional 8 h. After washing with cold Bio Legend cell staining buffer, cells were pelleted by centrifugation, resuspended in Annexin V binding buffer containing Annexin V‐APC and propidium iodide (PI), and gently vortexed to ensure uniform mixing. The mixture was incubated in the dark for 30 min and immediately subjected to flow cytometric analysis. Data were acquired and analyzed using FlowJo 10.8.1 software.

5.6. Gel Contraction Assay of hUSMCs

hUSMCs were suspended in rat tail type I collagen at an appropriate concentration and seeded into 24‐well plates. Following gel polymerization, DMEM containing TPT NP (100, 200, and 400 µg/mL) or atosiban (0.27 µg/mL) was added to the gels. After 24 h of incubation, fresh medium supplemented with LPS (1 µg/mL) was introduced, and the gels were cultured at 37°C for an additional 24 or 48 h. Untreated hUSMCs displayed intrinsic contractile activity, resulting in a measurable reduction in gel surface area over time. Gel images were acquired at 0, 24, and 48 h using a Canoscan 9000F scanner. The surface areas were measured with ImageJ software, and the values corresponding to 24 and 48 h were expressed as percentages of the 0 h baseline area.

5.7. Animals

Female Institute for Cancer Research (ICR) mice (8 weeks, 25–28 g) and male ICR mice (8 weeks, 25–28 g) were obtained from the Hunan Slake Experimental Animals Limited Company. Mice were raised under standard conditions with free feeding and drinking. Male and female mice were mated overnight, and pregnant mice were confirmed by positive vaginal smears the next day. Gestational day 0 and gestational period were defined as E0.5 and E19.5, respectively. This study established a PTB model in pregnant mice though intrauterine LPS‐induced inflammation, followed by different interventions. Comprehensive standardized husbandry was implemented throughout the experimental timeline, encompassing strict environmental controls, dietary assurance, and stress minimization protocols. Following different treatments, animals were euthanized, and biological samples including maternal blood, uterine tissues, and placentas were collected, with subsequent isolation of fetal tissues and organs for survival assessment and processing. All experimental procedures strictly complied with the Regulations on the Management of Laboratory Animals (China), the International Guiding Principles for Biomedical Research Involving Animals, and the OECD Guideline for the Testing of Chemicals: Prenatal Developmental Toxicity Study. All procedures and protocols were approved by the Ethics Committee of the First Affiliated Hospital of Chongqing Medical University (No. 2019–135). During the entire experiment, pregnant mice underwent daily monitoring of behavioral status, feeding patterns, and parturition indicators (such as restlessness and uterine contraction frequency), with intensified observation during anticipated therapy‐effect peaks. Fetal viability was assessed through continuous intrauterine activity monitoring and immediate postnatal evaluation. Humane endpoints were rigorously applied, requiring prompt euthanasia in accordance with experimental ethics guidelines when animals exhibited severe distress, moribund conditions, or irreversible experimental burden unresponsive to mitigation measures.

5.8. Establishment in a Mouse Model of PTB Induced by LPS

On embryonic day 16.5 (E16.5), pregnant mice were anesthetized with isoflurane. After induction of anesthesia, the abdominal region was disinfected, and a small laparotomy was performed. A 1.5 cm midline incision was made in the lower abdomen to expose the uterus. PTB was induced by injecting 20 µg LPS in 100 µL of saline between the first and second amniotic sacs in the lower portion of the right uterine horn of each mouse, while control mice received an equivalent volume of saline alone. Following the injection, the abdominal incision was closed using standard surgical procedures.

5.9. Tissue and Cellular Distribution Characteristics of TPT NP in Pregnant Mice

Pregnant mice with LPS‐induced PTB were randomly allocated into two groups: one received Cy5‐TPT NP via intravenous (i.v.) injection, and the other received an equivalent volume of saline as a control. At predetermined time points (0, 1, 2, 4, 8, 12, and 24 h), in vivo fluorescence imaging was performed for mice, and fluorescence intensities were quantified. Then nice were euthanized, and the uterus and placenta tissues were dissected for ex vivo fluorescence imaging. Both in vivo and ex vivo imaging were performed using an OI600 MF Touch multifunctional imaging system (BIO‐OI Biotechnology Co., Ltd., China). At 24 h after treatment, major maternal organs (heart, liver, spleen, lung, and kidney) and fetuses were harvested for ex vivo imaging, and fluorescence intensities of these samples were quantified. In addition, placentas collected at 24 h post‐treatment were embedded in OCT compound, and 7 µm cryosections were prepared and examined by fluorescence microscopy to acquire representative images.

5.10. Therapeutic Effects of TPT NP in Mice With LPS‐Induced Preterm Birth

To examine the therapeutic effects of TPT NP, preterm birth in pregnant mice was induced according to the previously described method. Subsequently, the pregnant mice were randomly assigned to different groups and subjected to different interventions. The control group received an intrauterine injection of saline solution, while the model group received an intrauterine injection of 20 µg LPS per mouse. After 30 min of intrauterine injection of 20 µg LPS, the TPT NP groups were treated by i.v. injection of different doses of TPT NP (2, 5, and 10 mg/kg), while the atosiban group received i.v. injection of atosiban at 0.17 mg/kg. Both the control and model groups received i.v. injection of an equal volume of saline solution.

PTB is defined as delivery occurring within 48 h after LPS induction. The pregnancy rates of pregnant mice in each group were calculated, and the numbers of full‐term and preterm pups were recorded. The percentage of surviving fetuses was calculated based on the total number of fetuses. After delivery, fetuses and placentas were collected, and macroscopic photographs were taken along with weight measurement. Also, the crown‐rump length (CRL) of fetuses was analyzed. For each litter, placental tissues and half of the fetuses were fixed in 4% paraformaldehyde, followed by hematoxylin and eosin (H&E) staining. Through histological observation, placental and fetal morphological changes were assessed. In addition, the distance ratio of spongiotrophoblast layer/labyrinth (S/L) and the maximal invasion distance of placenta were quantitatively analyzed. The remaining half of the fetuses were preserved in 95% ethanol, underwent organ enucleation, and subsequently processed for skeletal morphological assessment using Alcian blue and Alizarin red staining. In another parallel experiment, pregnant mice were randomly divided into the following groups: control, model, TPT NP (10 mg/kg), atosiban (0.17 mg/kg), and indomethacin (1 mg/kg). All interventions were administered via i.v. injection. The pregnancy rates in each group were then calculated.

5.11. Studies on Mechanisms Underlying Therapeutic Effects of TPT NP in PTB Mice

Similarly, PTB was defined as delivery occurring within 48 h after LPS injection, which induced PTB in nearly 100% of pregnant mice. At 8 h after LPS injection, pregnant mice were euthanized, and uterine, placental, and fetal membrane tissues were collected for analysis. Briefly, frozen sections of these tissues were prepared, followed by DHE staining for ROS assessment and MPO staining for neutrophil infiltration, both of which were imaged and quantitatively analyzed. Meanwhile, Cx43 immunofluorescence staining or Western blot analysis to analyze uterine contraction, mediators. Additionally, immunohistochemical analysis was performed to detect IL‐6 expression in uterine tissues. The mRNA expression levels of TNF‐α, IL‐1β, and IL‐6 in uterine, placental, and fetal membrane tissues were detected by qPCR. The primers used for mouse tissues were as follows: TNF‐α (forward, ACGGCATGGATCTCAAAGACA; reverse, TGAGAGCACGTAGTCGG), IL‐1β (forward, CAACTGCACTACAGGCTCCG; reverse, GTGGGTGTCCGTCTTTCAT), IL‐6 (forward, TGCTCTGGTCTTCTGGAGTTC; reverse, TGGAAGTTGGGGTAGGAAGGA), and GAPDH (forward, CAGTGGCAAAGTGGAGATTGTTG; reverse, TCGCTCCTGGAAGATGGTGAT). Moreover, the levels of H₂O₂, MDA, SOD, GPx, CAT, and TEAC in the above‐mentioned tissues were quantified by commercial kits. Additionally, peripheral blood was collected from mice in different groups, and WBC counts were measured to comprehensively evaluate the anti‐inflammatory efficacy of TPT NP in vivo.

Doppler ultrasound examination (Vevo 3100 LT, FujiFilm VisualSonics Inc.) was used to detect blood flow in the uterine artery and fetal umbilical artery after mice were exposed to LPS for 8 h. PTB mice after different interventions were anesthetized by inhalation of 5% isoflurane and oxygen (3 L/min flow rate), and anesthesia was maintained with 1.5% isoflurane in oxygen (2 L/min flow rate). Uterine artery doppler waveforms were collected from the branching point of the main trunk from the internal iliac artery, while fetal umbilical artery doppler waveforms were obtained from the free segment of the umbilical artery. By analyzing the peak systolic velocity (PSV), end‐diastolic velocity (EDV), and mean velocity (Vm) of the uterine artery and fetal umbilical artery, the therapeutic efficacy of TPT NP was evaluated. In addition, the resistance index (RI), pulse index (PI), and systolic/diastolic flow velocity ratio (S/D) were calculated according to the following formula: RI = (S − D)/S, PI = (S − D)/Vm, and S/D = PSV/EDV.

In a separate study, the PTB model was established in pregnant mice at embryonic day 16.5 (E16.5) by intrauterine injection of 20 µg LPS in each mouse. The control group received an equal volume of saline via the same route. At 30 min after LPS injection, the TPT NP groups were treated by i.v. injection of TPT NP at 2, 5, and 10 mg/kg. The atosiban group received an i.v. injection of atosiban at 0.17 mg/kg, while both the control and model groups were given an equivalent volume of saline. At 8 h after LPS exposure, mice were anesthetized, and a midline abdominal incision was made to expose the uterus. Throughout the procedure, care was exercised to minimize excessive traction and prevent vascular injury, thereby preserving the physiological integrity of the uterus. The uterine tissue was attached directly to a force transducer (RM6240E, Sichuan, China), and signals were transmitted to a computer via a signal converter. Data acquisition and recording were performed using the BL‐420F Biofunctional Experimental System software. The system recorded spontaneous uterine contraction tension over time and calculated parameters including mean muscle tension, maximum muscle tension, and area under the curve (AUC) to assess uterine contractility.

5.12. Statistical Analysis

Statistical analyses were performed using SPSS 23.0 software. Flow cytometry data were analyzed with FlowJo 10.8.1. All quantitative data were presented as mean ± standard deviation (s.d.). Levene's test was conducted to ensure the homogeneity of variances. Data in accordance with normal distribution was compared by one‐way ANOVA with post hoc LSD test or Tamhane's T2 test for more than 2 groups.

Author Contributions

Y.G.T., Y.F.C., and H.M.Z. contributed equally to this work. Y.G.T.: methodology, investigation, validation, data curation, formal analysis, and visualization. Y.F.C.: methodology, investigation, validation, and data curation. H.M.Z.: methodology, investigation, and validation. H.M.W.: methodology, investigation, data curation, and formal analysis. Y.Y.M.: investigation and data curation. J.X.Z.: conceptualization, funding acquisition, project administration, supervision, and writing – review & editing. H.B.Q.: resources, funding acquisition, project administration, supervision, and writing – review & editing. J.C.: conceptualization, methodology, investigation, data analysis, writing – original draft, funding acquisition, project administration, supervision, and writing – review & editing.

Conflicts of Interest

The authors declare no conflict of interest.

Supporting information

Supporting File: adhm71198‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.