Plasmonically Reinforced Self‐Sufficient Nanozymes Dysregulating Redox Homeostasis for Augmented Cascade Catalytic Oncotherapy

ABSTRACT

Tumor nanocatalytic therapy represents a promising transformative technology for treating malignant tumors, yet is often limited by insufficient catalytic activity and adaptive tumor microenvironment (TME) resistance. Herein, we develop a plasmonic Pt‐CuO₂ nanozyme that integrates plasmonic enhancement with self‐sufficient H₂O₂ supply for augmented cascade catalytic therapy. The nanozyme comprises Pt nanoparticles (NPs) and CuO₂ nanodots co‐deposited on Au nanostars and encapsulated within PEGylated ZIF‐8 metal–organic frameworks (MOFs). It exhibits strong near‐infrared localized surface plasmon resonance, enables intrinsic H₂O₂ generation, and depletes glutathione, effectively disrupting intratumoral redox homeostasis. Under acidic TME conditions, the MOFs degrade, releasing Pt NPs and CuO₂ nanodots to produce ∙OH via peroxidase‐like and Fenton‐like catalytic reactions. Plasmonic heating and “hot electron” injection under 808 nm laser irradiation further promote ∙OH generation. This process induces mitochondrial dysfunction, suppresses adenosine triphosphate biosynthesis and downregulates heat shock proteins, thereby increasing thermal sensitivity of cancer cells and enhancing the efficacy of mild plasmonic hyperthermia therapy (PHT). In a triple‐negative breast cancer murine model, the nanozyme demonstrates superior anticancer performance through the synergy of plasmon‐enhanced catalysis, self‐supplied H₂O₂, redox homeostasis disruption, and mild PHT. This study provides a novel strategy for efficient nanocatalytic therapy with substantial potential for clinical translation.

A plasmonically reinforced self‐sufficient nanozyme, designed as the plasmonic Pt‐CuO₂ nanozyme, is reported for plasmon‐enhanced catalytic cancer therapy. This nanozyme integrates strong localized surface plasmon resonance, self‐supply H₂O₂ capability, and glutathione depletion functional, enabling highly efficient ∙OH generation via peroxidase‐like and Fenton‐like reactions. In vivo studies demonstrate exceptional anticancer performance against triple‐negative breast cancer in mice.

1. Introduction

The marriage of nanomedicine with catalytic technology has spurred the emergence of nanocatalytic medicine, an evolving therapeutic paradigm aimed at the targeted treatment of malignant tumors [1, 2, 3]. Central to this approach is the use of biocompatible nanocatalysts to generate toxic reactive oxygen species (ROS) such as hydroxyl radicals (·OH), which trigger programmed apoptosis of cancer cells. Crucially, the tumor microenvironment (TME) presents distinctive biochemical features—including mild acidity, hypoxia, elevated glutathione (GSH) levels (1–10 mm), and excessive H₂O₂ production (50–100 µm), which differ fundamentally from those of healthy tissues [4, 5, 6, 7, 8, 9]. In nanocatalytic therapy, nanocatalysts convert overexpressed H₂O₂ within the TME into highly cytotoxic ·OH, enabling selective cancer cell destruction [10]. Key advantages of nanocatalysts over natural enzymes include flexible design, customizable enzymatic activities, efficient tumor accumulation, high stability, and cost‐effectiveness. These attributes make nanocatalytic platforms highly attractive for oncotherapy. Consequently, substantial research efforts have been dedicated to developing versatile nanocatalysts for anticancer treatments, with nanozymes and Fenton or Fenton‐like agents representing the most prominent candidates [11, 12, 13, 14, 15, 16]. Despite considerable advances, the clinical translation of nanocatalytic therapy remains challenging due to limited therapeutic efficacy, which arises from several interrelated factors: [6, 17, 18] (1) inefficient single‐enzyme activity of conventional nanocatalysts, (2) suboptimal catalytic kinetics, (3) constrained intratumoral H₂O₂ supply, and (4) a potent antioxidant defense system that maintains redox homeostasis in cancer cells. Thus, there is a pressing need to develop alternative strategies to achieve highly efficient nanocatalytic therapy for cancer.

Integrating nanocatalysts with plasmonic noble‐metal nanostructures (e.g., Au, Ag) represents a promising strategy to enhance the therapeutic efficacy of catalytic oncotherapy [19, 20, 21]. These nanostructures exhibit exceptional localized surface plasmon resonance (LSPR) properties, characterized by optical extinction cross‐sections 4–5 orders‐of‐magnitude larger than those of conventional organic or inorganic semiconductors [22, 23]. The LSPR effect enhances catalytic performance through two interconnected pathways: plasmonic “hot electron” injection and localized photothermal conversion [24, 25]. First, photoexcited LSPR generates high‐energy “hot electrons” via Landau damping, which inject into adjacent catalytic sites (e.g., semiconductors, reactive interfaces) [25, 26]. These “hot electrons” directly participate in redox reactions (e.g., H₂O₂ reduction, ·OH generation), significantly lowering activation barriers and accelerating reaction kinetics – a mechanism widely validated in photocatalysis and photovoltaics. Second, non‐radiative decay of LSPR converts absorbed photons into localized thermal energy, elevating the temperature at catalytic sites [6, 27]. This thermal input enhances catalytic activity by increasing reactant collision frequency, reducing activation energy, and accelerating Arrehenius‐type reaction kinetics. Notably, cancer cells exhibit heightened sensitivity to mild hyperthermia (40 °C–45 °C) compared to normal cells [28, 29, 30]. By operating within this temperature window, mild plasmonic hyperthermia therapy (mild PHT) minimizes collateral tissue damage while synergizing with nanocatalytic therapy. The therapeutic efficacy of mild PHT can be further augmented by downregulating heat shock proteins (HSPs), thereby increasing cancer cell thermal sensitivity. Thus, plasmonic nanocatalysts achieve superior anticancer outcomes through tripartite synergy: (1) enhanced catalytic ROS generation via “hot electron” injection, (2) accelerated catalytic reaction kinetics via localized heating, and (3) disruption of cellular homeostasis and tumor sensitization to oxidative stress via integrated mild PHT [6, 27].

Another strategy to improve the therapeutic efficacy of nanocatalytic therapy involves selective disruption of redox homeostasis in cancer cells [31, 32]. When exposed to ·OH generated catalytically in the TME, cancer cells activate adaptive antioxidant mechanisms to maintain redox balance and mitigate oxidative damage. Overexpressed GSH in the TME serves as a key antioxidant that scavenges ROS, thereby diminishing ROS‐mediated cytotoxicity and compromising therapeutic outcomes [33]. Although nanocatalysts with high intrinsic activity can achieve effective cancer cell killing, efficient GSH depletion remains essential to disrupt adaptive redox homeostasis and enhance therapeutic effects. Additionally, although intracellular H₂O₂ levels are elevated in cancer cells compared to normal cells, sustained ·OH generation retains constrained by limited endogenous H₂O₂ supply [34, 35]. Strategies to increase intratumoral H₂O₂ levels, such as direct delivery of exogenous H₂O₂ or in situ activation of endogenous H₂O₂ production, have therefore been explored [36, 37, 38]. However, direct H₂O₂ delivery suffers from limitations including low delivery efficiency, premature leakage, and potential adverse effects. In situ H₂O₂ production via glucose oxidase consumes intratumoral O₂, which is further limited by tumor hypoxia. Recently, metal peroxides (e.g, MgO₂, CaO₂, CuO₂) have emerged as alternative H₂O₂ sources that respond to acidic TME conditions [39, 40]. Although metal peroxides are unstable during systemic circulation and prone to rapid decomposition, appropriate nanocarriers (e.g., mesoporous SiO₂, metal–organic frameworks (MOFs), liposomes) can improve their stability during in vivo delivery and promote effective tumor accumulation. Thus, developing strategies for sustained ROS production and efficient GSH depletion, enabled by precise delivery of metal peroxides, is critical for disrupting redox homeostasis and potentiating nanocatalytic anticancer efficacy.

Herein, we develop a plasmonically reinforced self‐sufficient nanozyme – termed plasmonic Pt‐CuO₂ nanozymes – that enable efficient catalytic ·OH generation, GSH depletion, and H₂O₂ self‐supply within cancer cells for plasmon‐enhanced cascade catalytic oncotherapy. The nanozyme is engineered as Pt/CuO₂ co‐decorated Au nanostars (AuSts) encapsulated within PEGylated ZIF‐8 MOFs (Scheme 1a). The therapeutic mechanism involves multiple steps (Scheme 1b). (1) After cellular internalization, the MOF layer undergoes rapid degradation in the acidic TME, releasing Pt nanoparticles (NPs) and CuO₂ nanodots for direct catalytic action. (2) CuO₂ nanodots react with protons to generate H₂O₂ in situ, while the released Cu²⁺ depletes GSH, disrupting redox homeostasis and potentiating nanocatalytic therapy. (3) The exposed Pt‐CuO₂ nanozymes catalyze H₂O₂ decomposition via peroxidase‐like (POD‐like) and Fenton‐like reactions, producing cytotoxic ·OH for cancer cell elimination. (4) Strong near‐infrared LSPR properties from AuSts induce localized photothermal effects and generate high‐density plasmonic “hot electrons”, which accelerate catalytic reaction kinetics and enhance therapeutic efficacy. (5) The nanozymes downregulate HSP expression in tumor cells by triggering targeted mitochondrial damage and inhibiting adenosine triphosphate (ATP) biosynthesis, thereby amplifying mild PHT efficacy at temperatures below 45 °C and minimizing off‐targeted tissue damage. In vivo studies demonstrate significant tumor suppression in a triple‐negative breast cancer (TNBC) murine model following intravenous administration (Scheme 1c). This therapeutic outcome stems from the synergistic integration of plasmon‐enhanced nanocatalytic therapy and mild PHT, augmented by redox homeostasis dysregulation and self‐sufficient H₂O₂ supply.

SCHEME 1.

Schematic illustration of the preparation and therapeutic mechanism of the plasmonic Pt‐CuO₂ nanozyme for plasmon‐enhanced cascade catalytic oncotherapy. (a) Preparation process of the plasmonic Pt‐CuO₂ nanozyme, involving the deposition of Pt nanoparticles and CuO₂ nanodots onto plasmonic AuSts, ZIF‐8 MOF encapsulation, and surface PEGylation. (b) Integrated multifunctional mechanisms of the nanozyme, including H₂O₂ self‐supply, GSH depletion, Pt/Cu⁺‐mediated catalytic ∙OH generation, plasmonic heating effect, and “hot electron” injection. (c) Therapeutic process of the plasmonic Pt‐CuO₂ nanozyme for plasmon‐enhanced cascade catalytic therapy. After tumor accumulation, the acidic TME rapidly degrades the ZIF‐8 MOFs, releasing Pt nanoparticles and CuO₂ nanodots. The CuO₂ nanodots react with protons to produce H₂O₂ and Cu²⁺, which depletes GSH with the formation of the Fenton‐like agent Cu⁺; subsequently, Pt nanoparticles and Cu⁺ ions catalyze H₂O₂ conversion into ∙OH for cancer cell killing, which downregulates HSPs and synergizes with mild PHT and “hot electron” injection to achieve enhanced cascade catalytic oncotherapy.

2. Results and Discussion

2.1. Fabrication and Characterization of the Plasmonic Pt‐CuO₂ Nanozyme

In this study, AuSts were selected as the plasmonic substrate due to their excellent LSPR properties, which arise from the high‐density “built‐in” hotspots at their protruding spikes. These AuSts were synthesized via a seed‐mediated method well‐established in our laboratory [41, 42]. The plasmonic Pt‐CuO₂ nanozyme is composed of Pt NPs and CuO₂ nanodots co‐deposited onto plasmonic AuSts, encapsulated within PEGylated ZIF‐8 MOFs. Under the acidic TME, ZIF‐8 MOFs undergo rapid degradation, releasing Pt NPs and CuO₂ nanodots. The CuO₂ nanodots compensate for H₂O₂ consumption in POD‐like and Fenton‐like reactions, promoting ·OH generation. Additionally, the released CuO₂ nanodots deplete intratumoral GSH, while the LSPR effect from AuSts enhances catalytic ·OH production through plasmonic heating and “hot electron” injection [22, 24]. The synthesis of the plasmonic Pt‐CuO₂ nanozyme involves a three‐step process, outlined in Scheme 1a. First, Pt NPs were deposited onto AuSts via wet‐chemistry reduction of H₂PtCl₆ using L‐ascorbic acid (AA), yielding Pt‐AuSts (Figure S1a) [40, 41, 42]. The lattice fringes of Pt NPs are distinctly visible on the AuSt surface, confirming the successful synthesis of the Pt‐AuSts (Figure S2). Then, pre‐synthesized CuO₂ nanodots were mixed with Pt‐AuSts (Figure S1(b,c)), followed by the addition of Zn(NO₃)₂·6H₂O and 2‐methylimidazole (2‐mIM) to form ZIF‐8 MOFs, resulting in the formation of Pt‐CuO₂@AuSt@MOFs (denoted as plasmonic Pt‐CuO₂@MOFs). Finally, PEGylation with 1, 2‐distearoyl‐sn‐glycero‐3‐phosphoethanolamine‐N‐(methoxy(polyethyleneglycol)‐2000) (DSPE‐mPEG) was performed to obtain PEGylated plasmonic Pt‐CuO₂@MOFs (denoted as the plasmonic Pt‐CuO₂ nanozyme) for improved systemic circulation and biocompatibility [6, 44].

The morphology and structural characteristics of the plasmonic Pt‐CuO₂ nanozyme were characterized. Transmission electron microscopy (TEM) and scanning electron microscopy (SEM) images demonstrate that the plasmonic Pt‐CuO₂ nanozyme exhibits a monodisperse core/shell structure with an average diameter of ≈176 nm (Figure 1(a,b) and Figure S1d). Energy‐dispersive X‐ray spectroscopy (EDS) elemental mapping shows that Pt and Cu elements are mainly concentrated in the AuSt core region, while Zn uniformly surrounds the structure, verifying the successful deposition of Pt NPs and CuO₂ nanodots on AuSts encapsulated within ZIF‐8 MOFs (Figure 1c). The powder X‐ray diffraction (XRD) analysis reveals distinct diffraction peaks corresponding to crystalline ZIF‐8 MOFs, and AuSts, while the amorphous structure of CuO₂ nanodots is evident (Figure 1d). The diffraction peaks corresponding to Pt NPs are not discernible in the XRD pattern of the Pt‐AuSts, which can be attributed to their relatively small particle size and the dominant diffraction pattern of the underlying AuSts. Fourier transform infrared (FTIR) spectroscopy further confirms the structural composition, displaying characteristic peaks at 1580 cm⁻¹ (─C═N‐ stretching of ZIF‐8 MOFs) and at 1109/1736 cm⁻¹ (P‐O asymmetric vibration and ─C═O stretching of DSPE‐mPEG), respectively (Figure S3) [6, 45, 46]. Dynamic light scattering measurements show that the hydrodynamic diameter is 50.3 (±3.0) nm for AuSts and 47.7 (±3.6) nm for Pt‐AuSts, but increases to 189.9 (±18.9) nm for plasmonic Pt‐CuO₂@MOFs and 198.0 (±11.4) nm for the plasmonic Pt‐CuO₂ nanozyme (Figure S1(e, i)). The ζ‐potentials of AuSts, Pt‐AuSts, Pt‐CuO₂@AuSt@MOFs, and the plasmonic Pt‐CuO₂ nanozyme are −20.0 (±1.2), ‐22.6 (±2.7), +15.4 (±1.27), and ‐11.3 (±1.1) mV, respectively (Figure S1(e, ii)). This again confirms successful DSPE‐mPEG modification of Pt‐CuO₂@AuSt@MOFs, which reverses the surface charge from positive to negative and thereby improves biocompatibility and circulation stability. We also prepared PEGylated AuSt@MOFs (denoted as AuSt‐PEG) and PEGylated Pt‐AuSt@MOFs (denoted as plasmonic Pt nanozyme) using a similar synthetic protocol, but in the absence of Pt NPs/CuO₂ nanodots and CuO₂ nanodots, respectively. The plasmonic Pt‐CuO₂ nanozyme demonstrates excellent structural stability in both PBS at pH 7.4 and cell culture medium (RPMI‐1640 supplemented with 10% fetal bovine serum (FBS)). This is evidenced by the absence of significant changes in both ζ‐potential and hydrodynamic size over a 7‐day incubation period (Figure S4). The loading capacity and encapsulation efficiency are quantified by an inductively coupled plasma‐optical emission spectrometer, yielding values of 90.3% and 5.1% for Pt, and 63.5% and 3.2% for CuO₂, respectively.

FIGURE 1.

Structural and photophysical characterization of the plasmonic Pt‐CuO₂ nanozyme. (a) TEM image, (b) SEM image, and (c) HAADF‐STEM image with the corresponding elemental maps of the plasmonic Pt‐CuO₂ nanozyme. (d) Powder XRD patterns of AuSts, CuO₂ nanodots, Pt‐AuSts, and the plasmonic Pt‐CuO₂ nanozyme. Blue and yellow vertical lines denote the reference diffraction peaks of Au (JCPDS #: 04–0784) and Pt (JCPDS #: 04–0802), respectively, and black vertical lines represent the simulated XRD pattern of ZIF‐8 MOFs. (e) Optical extinction spectra of AuSts, Pt‐AuSts, and the plasmonic Pt‐CuO₂ nanozyme dispersed in water. (f) 3D distributions of (i) near‐field enhancement (log₁₀(|E|/|E₀|)²) and (ii) temperature increase (ΔT, °C) excited at 808 nm. The maximum near‐field enhancement value and the range of the temperature rise (ΔT_min, ΔT_max) are indicated at the bottom. (g) Infrared thermal images of aqueous suspensions containing different concentration of the plasmonic Pt‐CuO₂ nanozyme (0 (PBS), 25, 50, 125, and 250 µg/mL) irradiated with an 808 nm laser (1.0 W/cm²) at varying durations. (h) (i) The temperature evolution curves of nanozyme suspensions at different concentrations under laser irradiation; (ii) photothermal heating‐cooling profile and corresponding Time versus ‐ln(θ) plot during the cooling phase for the nanozyme suspension at 250 µg/mL.

Next, optical extinction spectroscopy was employed to monitor the synthesis process of the plasmonic Pt‐CuO₂ nanozyme. As shown in Figure 1e, the as‐synthesized AuSts display a LSPR band at 775 nm. After Pt NP deposition, the LSPR band broadens with weakened plasmon strength and no obvious spectral shift, which is attributed to plasmon damping at the Au‐Pt interface [41]. Subsequent encapsulation with ZIF‐8 MOFs results in a red shift of the LSPR peak to 800 nm in the final plasmonic Pt‐CuO₂ nanozyme, owing to the increased local refractive index [42]. We further conducted a theoretical analysis of the plasmonic and photothermal properties using the finite‐element method in COMSOL Multiphysics 5.6. The AuSt was modeled as an Au nanosphere with ten uniformly distributed spikes on its three‐dimensional (3D) surface, and the Pt‐AuSt was constructed by attaching seven Pt semi‐spheres to each spike of the AuSt; detailed dimensional parameters are provided in Figure S5(a,b). The simulated extinction cross‐section spectra reveal a prominent LSPR band at 790 nm for AuSt and 786 nm for Pt‐AuSt (Figure S5c). We computed the 3D distribution of near‐field enhancement and the plasmonic heating‐induced temperature increase (ΔT) at 808 nm for both AuSts and Pt‐AuSts, a commonly used wavelength for near‐infrared photothermal therapy. As illustrated in Figure 1f, both structures exhibit maximum near‐field enhancement at the spike tips and minimum enhancement on the core. In contrast, the temperature rise distribution is highly uniform across the entire structure, with a variation of approximately 1 °C [22]. Notably, the deposition of Pt on AuSts leads to a slight reduction in both near‐field enhancement and photothermal temperature elevation. These results confirm that both AuSts and Pt‐AuSts possess excellent near‐infrared plasmonic properties and effective photothermal performance.

The excellent near‐infrared plasmonic properties of the plasmonic Pt‐CuO₂ nanozyme suggest its potential as a photothermal agent for PHT. We therefore evaluated its photothermal performance under 808 nm laser irradiation at a power density of 1.0 W/cm². As shown in Figure 1g, the temperature of the plasmonic Pt‐CuO₂ nanozyme aqueous suspensions increases gradually with prolonged laser irradiation, exhibiting a clear concentration‐dependent temperature rise. After 10 min of irradiation, the suspension at 250 µg/mL reaches a steady‐state temperature of 55.5 °C, whereas pure water under the same conditions only reaches 28.6 °C (Figure 1(h, i)). The temperature rise also correlates positively with laser power density ranging from 0–1.0 W/cm² investigated in this work (Figure S6a). The photothermal conversion efficiency of the plasmonic Pt‐CuO₂ nanozyme is determined to be 43.3% based on the heating‐cooling curves using a well‐established method (Figure 1(h, ii)). This value is comparable to, or slightly lower than, those reported for pristine spiky Au nanostructures (Table S1). The observed decrease can be attributed to the weakened plasmonic strength resulting from the deposition of Pt/CuO₂ and ZIF‐8 coating on AuSts, as evidenced in Figure 1e. It is important to note that the intended application in this work is mild PHT (40 °C–45 °C), for which the achieved photothermal performance remains fully sufficient. Over four laser on/off cycles, no significant decrease in the steady‐state temperature is observed, indicating excellent photostability (Figure S6b). These results demonstrate that the plasmonic Pt‐CuO₂ nanozyme can function as a reliable photothermal agent for in vivo mild PHT through adjusting both nanozyme concentration and laser power density.

2.2. Plasmon‐Enhanced Cascade Catalytic Performance

The plasmonic Pt‐CuO₂ nanozyme integrates multiple therapeutic functionalities, including H₂O₂ self‐supply, GSH depletion, Pt‐based POD‐like activity and Cu⁺‐mediated Fenton‐like activity for catalytic ·OH generation, plasmonic heating effects, and plasmonic “hot electron” injection (Figure 2a). First, the degradation and payload release kinetics of the plasmonic Pt‐CuO₂ nanozyme were assessed under simulated physiological conditions in RPMI‐1640 medium at pH 7.4 and 5.4. As shown in Figure S7a, the nanozyme exhibits no significant morphological change after 3 h of incubation at pH 7.4, while its MOF layer was completely degraded at pH 5.4. Further incubation for 24 h reveals a cumulative Cu release of 72.4% at pH 5.4, significantly higher than that (5.6%) observed at 7.4 (Figure S7b). These results confirm the acidic TME‐triggered degradation of the MOF layer and the corresponding release of payloads. The ·OH generation was evaluated by electron spin resonance (ESR) spectroscopy with 5,5‐dimethyl‐1‐pyrroline‐N‐oxide (DMPO) as a trapping agent [49, 50]. As shown in Figure 2b, a characteristic quartet pattern with a peak intensity ratio of 1:2:2:1 is clearly observed in the ESR spectrum of the plasmonic Pt‐CuO₂ nanozyme in the dark, confirming ·OH generation from self‐supplied H₂O₂ by the POD‐like reaction under acidic conditions. The ESR signals of DMPO/·OH are enhanced upon external H₂O₂ addition, leading to increased ·OH production. Furthermore, laser irradiation amplifies the ESR signals of DMPO/·OH, demonstrating plasmonic enhancement of catalytic ·OH generation.

FIGURE 2.

In vitro evaluation of plasmon‐enhanced catalytic activity of the plasmonic Pt‐CuO₂ nanozyme. (a) Schematic illustration of the acidity‐responsive H₂O₂ self‐supply, plasmon‐enhanced catalytic ∙OH generation, and GSH depletion by the plasmonic Pt‐CuO₂ nanozyme, along with the corresponding detection methods. (b) ESR spectra of DMPO/∙OH adducts after treatment with the plasmonic Pt‐CuO₂ nanozyme in the presence or absence of 10 mm H₂O₂ either in the dark (‐) or under 808 nm laser irradiation (1.0 W/cm², 5 min). (c) (i) Absorption spectra of 10 mm TMB aqueous solutions (pH 5.4) treated with various concentrations (0, 10, 20, and 40 µg/mL) of the plasmonic Pt‐CuO₂ nanozyme in the presence of 10 mm H₂O₂, (ii) time‐dependent absorbance at 652 nm of TMB solution treated by 1.0 mg/mL plasmonic Pt‐CuO₂ nanozyme at varied pH values (pH 5.4, 6.5, and 7.4), and (iii) Michaelis–Menten kinetic analysis using H₂O₂ as the substrate, performed in the dark or under 808 nm laser (1.0 W/cm², 5 min). (d) (i) Absorption spectra of the TMB solution treated with 1.0 mg/mL plasmonic Pt nanozyme in the dark, under 808 nm laser (1.0 W/cm², 5 min), and under 808 nm laser with ice cooling, (ii) absorbance at 652 nm of TMB after treatment with 1.0 mg/mL plasmonic Pt nanozyme under 808 nm laser at various power densities (0‐500 mW/cm²) for 5 min, and (iii) absorbance at 652 nm of TMB treated with 0.5 mg/mL plasmonic Pt nanozyme in the presence or absence of ethanol (EtOH) in dark or under 808 nm laser (1.0 W/cm², 5 min). (e) (i) Absorption spectra of 0.316 mm KMnO₄ solutions treated with H₂O (control), 10 mm H₂O₂, 150 mm CuO₂, and 200 mm plasmonic Pt‐CuO₂ nanozyme, (ii) H₂O₂ production by the plasmonic Pt‐CuO₂ nanozyme at different concentrations, and (iii) relative GSH levels after treatment with 1.0 mg/mL plasmonic Pt‐CuO₂ nanozyme at various pH values (pH 5.4, 6.5, and 7.4) for different durations. Data are presented as mean ± standard deviation (n = 3).

The ·OH generation was further examined using the 3,3′,5,5′‐tetramethylbenzidine (TMB) colorimetric assay in the presence of H₂O₂ [11]. Oxidation of TMB by the highly reactive ·OH yields blue oxTMB, characterized by an absorption peak at 652 nm. As depicted in Figure 2(c, i), the absorbance at 652 nm increases progressively with rising concentrations of the plasmonic Pt‐CuO₂ nanozyme, indicating enhanced ·OH production under acidic conditions. The pH‐dependent ·OH generation was also investigated at pH 7.4, 6.5, and 5.4, respectively (Figure 2(c, ii)). At pH 7.4, no significant change in oxTMB absorbance is detected, suggesting minimal release of Pt NPs and CuO₂ nanodots and consequently negligible ·OH generation under physiological conditions. The absorbance at 652 nm increases slightly over time at pH 6.5, and markedly at pH 5.4, confirming the highest catalytic activity of the nanozyme under acidic conditions. Steady‐state catalytic kinetics was evaluated by varying the H₂O₂ concentration in the dark or under 808 nm laser irradiation (1.0 W/cm²). The velocity (V) of ·OH formation was assessed based on TMB oxidation. The Michaelis‐Menten curves were fitted to the velocity versus H₂O₂ concentration data (Figure 2(c, iii)), and key parameters were derived from Lineweaver–Burk plots (Figure S8). The maximum reaction velocity (V _max) and Michaelis–Menten constant (K _m) are 11.2 nm s⁻¹ and 8.17 mm with no laser irradiation, and 14.0 nm s⁻¹ and 8.08 mm with laser irradiation, respectively. These results indicate that laser irradiation significantly enhances the catalytic efficiency and substrate affinity of the nanozyme for ·OH generation, much superior to those nanozymes previously reported in the literature (Table S2). A careful analysis reveals that our nanozyme exhibits a strong affinity for H₂O₂, as indicated by its lower K _m compared to non‐Pt‐based metallic nanozymes. This ensures efficient catalytic activity even at the low endogenous concentrations found in the TME. It is noteworthy that our nanozyme achieves a relatively lower V _max while maintaining high atomic utilization efficiency, particularly when compared to nanozymes composed of pure Pt or those with high Pt loadings.

The contribution of the LSPR effect to catalytic enhancement was further elucidated. Plasmonic mechanisms primarily include plasmonic heating and “hot electron” injection. We observed a pronounced photocurrent response from the plasmonic Pt‐CuO₂ nanozyme‐based working electrode under 808 nm laser irradiation (Figure S9), confirming the photo‐induced injection of “hot electrons”. Using Pt‐AuSts as a model system, ·OH generation was assessed in the presence of H₂O₂ and TMB. As shown in Figure 2(d, i), laser irradiation markedly increases the oxTMB absorbance at 652 nm compared to the dark condition, underscoring the crucial role of LSPR in enhancing catalytic performance. To decouple the effects of plasmonic heating and “hot electron” injection, experiments were conducted under ice cooling during laser irradiation to maintain a constant temperature at room temperature, thereby eliminating thermal contributions. The results show that the oxTMB absorbance decreases significantly under ice cooling yet remains higher than that without laser, suggesting that both plasmonic heating and “hot electron” injection contribute to the enhanced ·OH generation with the latter playing a more dominant role. Additionally, the influence of laser power density on ·OH generation was investigated. As illustrated in Figure 2(d, ii), the oxTMB absorbance rises noticeably with increasing laser power density from 0 to 500 mW/cm², indicating enhanced ·OH production at higher power densities. The effect of ethanol, which promotes electron‐hole separation by acting as a hole scavenger, was also examined (Figure 2(d, iii)). Introduction of ethanol under laser irradiation significantly boosts ·OH generation, further corroborating the involvement of “hot electron” injection in the plasmon‐enhanced catalytic process. In addition, we confirm that the CAT‐like activity of the plasmonic Pt‐CuO₂ nanozyme is significantly inhibited under acidic TME (Figure S10).

2.3. GSH Depletion and H₂O₂ Self‐Supply for Redox Homeostasis Dysregulation

Excessive ROS accumulation is conductive to amplifying oxidative stress in cancer cells. However, the TME exhibits insufficient endogenous H₂O₂ for ·OH production, and the elevated GSH levels consume the ·OH, compromising the efficacy of catalytic therapy [33, 34, 35]. In the present design, the plasmonic Pt‐CuO₂ nanozyme enables H₂O₂ self‐supply exclusively at tumor sites through the protonation of loaded CuO₂ nanodots, while depleting GSH via released Cu²⁺. The KMnO₄ assay was utilized to evaluate the H₂O₂ generation capacity of the plasmonic Pt‐CuO₂ nanozyme under an acidic environment [51]. KMnO₄ reacts with H₂O₂ to produce Mn²⁺ and O₂, accompanying by a color change from purple to colorless (Figure 2a). As shown in Figure 2(e, i), the characteristic absorption peak of KMnO₄ disappears in the absorption spectra of the plasmonic Pt‐CuO₂ nanozyme suspension, confirming H₂O₂ production from CuO₂ under acidic conditions (pH 5.4). The H₂O₂ production increases with elevated nanozyme concentrations at acidic pH (Figure 2(e, ii) and Figure S11). Given the oxidative property of released Cu²⁺, the GSH depletion capability was further investigated using 5,5'‐dithiobis‐(2‐nitrobenzoic acid) (DTNB) chromogenic assay [50]. As expected, the GSH level gradually decreases with prolonged incubation time in the presence of the nanozyme, with the highest GSH depletion rate observed at pH 5.4 due to enhanced Cu²⁺ release (Figure 2(e, iii)). The GSH‐mediated reduction of Cu²⁺ to Cu⁺ was further investigated using X‐ray photoelectron spectroscopy (XPS). Upon GSH treatment, the Cu²⁺/Cu⁺ molar ratio of the plasmonic Pt‐CuO₂ nanozyme decreased significantly from 1.88 to 0.4 (Figure S12), directly confirming the effective reduction of Cu²⁺ species. The results demonstrate that the plasmonic Pt‐CuO₂ nanozyme degrades in acidic environments, enabling H₂O₂ self‐supply, GSH depletion, and enhanced ·OH generation under laser irradiation.

2.4. In Vitro Anticancer Efficacy and Therapeutic Mechanisms

The anticancer efficacy of the plasmonic Pt‐CuO₂ nanozyme was evaluated in vitro using 4T1 TNBC cells and L‐02 human normal liver cells (Figure 3a). Biocompatibility assessment reveals that the cell viability of L‐02 cells remains high (>82%) at the nanozyme concentration up to 180 µg/mL, indicating favorable biocompatibility (Figure S13). In contrast, the cell viability of 4T1 cells decreases to 65% at the same concentration, attributable to specific ·OH generation within cancer cells. Comparative studies of AuSt‐PEG, plasmonic Pt nanozyme, and the plasmonic Pt‐CuO₂ nanozyme were conducted both in the dark (‐) and under 808 nm laser irradiation (+), demonstrating distinct anticancer effects. As shown in Figure 3b, AuSt‐PEG exhibits high biocompatibility, with cell viability exceeding 88% even at 180 µg/mL. In comparison, the plasmonic Pt nanozyme and the plasmonic Pt‐CuO₂ nanozyme reduce the cell viability of 4T1 cells to 66% and 57%, respectively, due to their enhanced catalytic ·OH generation and H₂O₂ self‐supply capacities. Laser irradiation (+) further reduces the cell viability to 77% for AuSt‐PEG, 26% for the plasmonic Pt nanozyme, and 12% for the plasmonic Pt‐CuO₂ nanozyme. The relatively high cell survival in the AuSt‐PEG (+) group may stem from HSP‐mediated thermal tolerance in cancer cells. Notably, the incorporation of CuO₂ in the plasmonic Pt‐CuO₂ nanozyme significantly augments cancer cell killing under laser irradiation compared to the plasmonic Pt nanozyme, owing to H₂O₂ self‐supply and Cu⁺‐based Fenton‐like reaction. Further, the synergistic effect among the therapeutic components was quantitatively evaluated based on the cytotoxicity data using the Chou‐Talalay method (Figure 3b and Figure S14) [47]. The calculated combination index (CI) for the combined treatment of plasmonic Pt nanozyme, AuSt‐PEG with laser, and CuO₂ nanodots is 0.83. According to the established criterion where a CI value less than 1.0 indicates synergism, this result confirms a synergistic interaction within our nanozyme system. These results underscore the superior anticancer activity of the plasmonic Pt‐CuO₂ nanozyme, resulting from the synergistic combination of PHT, H₂O₂ self‐supply, and plasmon‐enhanced catalytic ·OH generation. Consistent trends were also observed in the Calcein‐AM/propidium iodide (PI) co‐staining assay (Figure S15).

FIGURE 3.

In vitro evaluation of cytotoxicity, redox homeostasis disruption, and anticancer efficacy. (a) Schematic illustration of the anticancer mechanism involving plasmon‐enhanced cascade catalytic reactions and disruption of cellular redox homeostasis. (b) Cell viability of 4T1 cells treated with AuSt‐PEG, plasmonic Pt nanozyme, and plasmonic Pt‐CuO₂ nanozyme at varying concentrations for 24 h in the dark (‐) or under 808 nm laser irradiation (1.0 W/cm², 5 min) (+) (n = 5 per group). (c) Fluorescence images of 4T1 cells incubated with Rhodamine 6G‐labeled plasmonic Pt‐CuO₂ nanozyme for 0, 3, 6, and 9 h for cellular uptake tracking. (d) Intracellular ROS generation detected by DCFH‐DA staining in 4T1 cells receiving various treatments. (e) Relative intracellular GSH and ATP levels in 4T1 cells after 6 h treatment at different concentrations of plasmonic Pt‐CuO₂ nanozyme, normalized to the PBS‐treated control (n = 5 per group). (f) Alternations in mitochondrial membrane potential assessed by JC‐1 assay in 4T1 cells after 12 h of treatment with the indicated formulations. (g) Western blot analysis of Hsp70 and Hsp90 expression in 4T1 cells receiving various treatments. Data are presented as mean ± standard deviation (n = 5), and statistical analysis was carried out using one‐tail Student's t‐test (^**** p <0.0001).

Effective cellular internalization is critical for anticancer performance. Using Rhodamine 6G (Rh6G) labeling, we tracked the cellular uptake of the plasmonic Pt‐CuO₂ nanozyme in 4T1 cells. As shown in Figure 3c, intracellular red fluorescence intensifies over culture time, confirming efficient endocytosis of the plasmonic Pt‐CuO₂ nanozyme. We further probed intracellular ·OH generation using 2',7′‐dichlorofluorescein diacetate (DCFH‐DA) probe, which emits green fluorescence upon oxidation by ·OH. As illustrated in Figure 3d, negligible green fluorescence is detected in 4T1 cells treated with AuSt‐PEG either in the dark (‐) or under laser irradiation (1.0 W/cm²) (+). Weak fluorescence is observed in the plasmonic Pt nanozyme (‐) group, while a stronger fluorescence signal is evident in cells treated with the plasmonic Pt‐CuO₂ nanozyme (‐), due to Pt/Cu⁺‐based catalytic ∙OH generation augmented by H₂O₂ self‐supply. Moreover, laser irradiation markedly enhances green fluorescence in both plasmonic Pt nanozyme (+) and plasmonic Pt‐CuO₂ nanozyme (+) groups, with the latter exhibiting the strongest signal, confirming plasmon‐enhanced catalytic ·OH generation. Furthermore, we observe that the introduction of catalase significantly inhibits ∙OH generation in 4T1 cells treated with CuO₂ nanodots (Figure S16). This finding underscores the central role of H₂O₂ supply in ∙OH production, as catalase efficiently decomposes H₂O₂ into water and O₂, thereby depleting the essential substrate for the POD‐like and Fenton‐like reactions that generate the cytotoxic ∙OH species.

Despite efficient ·OH production via Pt/Cu⁺‐catalyzed reactions, high intracellular GSH levels in cancer cells can scavenge ·OH and maintain redox homeostasis, potentially compromising the therapeutic outcomes [33]. We thus evaluated the GSH depletion capability of the nanozyme in the cellular level. In our design, released Cu²⁺ under acidic conditions can directly oxidize GSH to GSSG. Treatment with 180 µg/mL plasmonic Pt‐CuO₂ nanozyme reduces intracellular GSH levels by 58%, demonstrating effective redox homeostasis disruption (Figure 3e). Given the role of mitochondria as the cellular energy supply sites and their susceptibility to ROS, we assessed the mitochondrial membrane potential (MMP) in 4T1 cells after various treatments using the JC‐1 assay. Healthy mitochondria with high MMP exhibit red JC‐1 aggregates, while apoptotic cells with diminished MMP show green monomeric fluorescence [30]. As shown in Figure 3f, 4T1 cells treated with PBS or AuSt‐PEG maintain strong red fluorescence regardless of laser exposure, indicating intact mitochondria. Cells treated with the plasmonic Pt‐CuO₂ nanozyme (‐) produce moderate green fluorescence, slightly higher than in the plasmonic Pt nanozyme (‐) group, consistent with enhanced ·OH generation. Under 808 nm laser irradiation, green fluorescence becomes dominant in both plasmonic Pt nanozyme (+) and plasmonic Pt‐CuO₂ nanozyme (+) groups, with the latter exhibiting the most severe mitochondrial damage. Considering that ATP is mainly produced in mitochondria, we further investigated the intracellular ATP level in 4T1 cells treated with varying concentrations of the plasmonic Pt‐CuO₂ nanozyme. It is shown that the intracellular ATP level decreases gradually with the elevated nanozyme concentration, declining by >50% at 180 µg/mL (Figure 3e). Since HSP expression depends on mitochondrial ATP supply, we performed Western blot analysis of Hsp70 and Hsp90. Both plasmonic Pt nanozyme (+) and plasmonic Pt‐CuO₂ nanozyme (+) treatments downregulate Hsp70 and Hsp90 expression in 4T1 cells, with the most substantial reduction observed in the plasmonic Pt‐CuO₂ nanozyme (+) group (Figure 3g). Furthermore, the downregulation of both HSP70 and HSP90 was significantly reversed in 4T1 cells treated with the plasmonic Pt‐CuO₂ nanozyme (+) upon the addition of GSH as an ROS scavenger (Figure S17). This confirms that the generated ROS are directly responsible for the suppression of HSP expression. Collectively, these results demonstrate that the plasmonic Pt‐CuO₂ nanozyme induces mitochondrial dysfunction and suppresses HSP expression via an ROS‐mediated pathway, thereby sensitizing the cancer cells to PHT.

2.5. In Vivo Antitumor Efficacy and Biosafety

Encouraged by the excellent cascade catalytic performance, the in vivo therapeutic efficacy of the plasmonic Pt‐CuO₂ nanozyme was further evaluated in 4T1 tumor‐bearing mice, following the experimental timeline illustrated in Figure 4a. Prior to in vivo anticancer assessment, we examined the biosafety, pharmacokinetics, and biodistribution of the plasmonic Pt‐CuO₂ nanozyme. Hemocompatibility was assessed by testing the hemolytic effect of the nanozyme on red blood cells (RBCs). After incubation with various concentrations (12.5–400 µg/mL), the hemolysis rate remains significantly below the acceptable threshold of 5% even at the highest concentration up to 400 µg/mL (Figure S18), demonstrating negligible hemolytic activity and good biocompatibility. For pharmacokinetics and biodistribution analysis, 4T1 tumor‐bearing mice were intravenously administrated with the plasmonic Pt‐CuO₂ nanozyme, and blood was collected at different time intervals for quantitative Au analysis via inductively coupled plasma‐mass spectrometry. The blood circulation profile follows a typical two‐compartment model with half‐lives of t_1/2α = 0.49 h and t_1/2β = 3.26 h, suggesting sufficient circulation time for tumor accumulation (inset, Figure 4b). Biodistribution studies in major organs (heart, liver, spleen, lung, and kidney) and tumors at 6, 12, and 24 h post‐administration reveal predominant accumulation in the liver, spleen, lung, and kidney due to reticuloendothelial system clearance. Notably, significant tumor accumulation (≈7.1% ID/g) was observed at 12 h post‐administration, attributable to the enhanced permeability and retention effect. These results confirm the favorable biosafety, prolonged blood circulation, and efficient tumor targeting of the nanozyme.

FIGURE 4.

In vivo evaluation of the anticancer efficacy of the plasmonic Pt‐CuO₂ nanozyme. (a) Schematic illustration of the experimental procedure for tumor model establishment and treatment protocol. (b) Biodistribution of Au across major organs and tumors at 6, 12, and 24 h post‐intravenous administration of the plasmonic Pt‐CuO₂ nanozyme (n = 3). Inset: corresponding blood circulation profile. (c) (i) Infrared thermal images and (ii) temperature elevation curves of tumor regions in 4T1 tumor‐bearing mice following intravenous administration of PBS or the plasmonic Pt‐CuO₂ nanozyme under 808 nm laser irradiation (1.0 W/cm²). (d) Representative photographs and (e) individual tumor volume growth curves of across different treatment groups: PBS + laser, AuSt‐PEG + laser, plasmonic Pt‐CuO₂ nanozyme, and plasmonic Pt‐CuO₂ nanozyme + laser (n = 5). Laser irradiation (808 nm, 1.0 W/cm², 5 min) was applied at 12 h post‐administration. (f) Tumor growth progression in each treatment group over the study period. (g) (i) Photographs and (ii) average weights of excised tumors collected on day 16. Data are presented as mean ± standard deviation, and statistical analysis was carried out using one‐tail Student's t‐test (^** p <0.01, ^*** p <0.001, and ^**** p <0.0001).

For in vivo anticancer efficacy evaluation, when the tumors reached 120 mm³, the 4T1 tumor‐bearing mice were randomly divided into four treatment groups (n = 5): (1) PBS + laser, (2) AuSt‐PEG + laser, (3) plasmonic Pt‐CuO₂ nanozyme, and (4) plasmonic Pt‐CuO₂ nanozyme + laser. Mice received intravenous administration of AuSt‐PEG or plasmonic Pt‐CuO₂ nanozyme at a dosage of 20 mg/kg on day 0. For laser irradiation (+) groups, the tumors were continuously irradiated with a 808 nm laser for 5 min at 12 h post‐administration. Tumor temperature was monitored in real‐time using a FLIR A35 infrared thermal camera. As presented in Figure 4c, the plasmonic Pt‐CuO₂ nanozyme (+) group reaches a steady‐state temperature of 44.9 °C, suitable for mild PHT, while the PBS (+) group achieves a steady‐state temperature of only 39 °C. Tumor volume and body weight were recorded every other day for 16 days. As shown in Figure 4(d–f), the PBS (+) group shows rapid tumor growth, while both AuSt‐PEG (+) and plasmonic Pt‐CuO₂ nanozyme (‐) groups exhibit significant tumor inhibition with 40.7% and 38.5% tumor inhibition rates, respectively. The plasmonic Pt‐CuO₂ nanozyme (+) group achieves the highest tumor suppression with a tumor inhibition rate of 73.8%, consistent with in vitro cytotoxicity results (Figure 4g). The superior anticancer efficacy is attributed to the synergistic combination of Pt/Cu⁺‐mediated catalytic therapy, H₂O₂ self‐supply, redox homeostasis disruption, and plasmonic enhancement. All groups maintained stable body weights during the course of treatments, indicating minimal systemic toxicity (Figure S19).

Further histological and immunohistochemical analyses were performed on excised tumors (Figure 5). Hematoxylin and eosin (H&E) staining reveals pronounced nuclear shrinkage in the plasmonic Pt‐CuO₂ nanozyme (+) group. DCFH‐DA staining of tumor sections revealed that the plasmonic Pt‐CuO₂ nanozyme (+) group exhibits the most intense ∙OH signal within the tumor tissue. The ∙OH generation observed in tumors treated with the plasmonic Pt‐CuO₂ nanozyme was significantly more effective than that achieved with AuSt‐PEG plus laser treatment. These results underscore the critical role of the CuO₂ nanodots in facilitating efficient ∙OH production within tumor tissues. Terminal deoxynucleotidyl transferase dUTP nick‐end labeling (TUNEL) staining shows the strongest green fluorescence in this group, indicating maximal cell apoptosis. Hsp70/Hsp90 immunofluorescence staining confirms significant downregulation of HSP expression, resulting from ROS‐induced mitochondrial damage and ATP supply blockade. Additionally, blood biochemical analysis of the mice after different treatments was conducted on day 16 post‐administration for alanine transaminase (ALT), alkaline phosphatase (ALP), blood urea nitrogen (BUN), aspartate transferase (AST), and lactate dehydrogenase (LDH). The levels of all these indicators fall within normal reference ranges, confirming negligible systemic toxicity (Figure S20). H&E staining of major organs reveals no obvious organ damage or abnormalities after 16 days of different treatments, further verifying the high biosafety of the nanozyme (Figure S21). Furthermore, after 30 days of the intravenous administration of the plasmonic Pt‐CuO₂ nanozyme, healthy mice exhibit no significant abnormalities in key blood biochemical indicators and show negligible pathological damage in major organs (Table S3 and Figure S22). These results demonstrate the excellent biocompatibility and long‐term biosafety of the plasmonic Pt‐CuO₂ nanozyme treatment.

FIGURE 5.

Histopathological and immunofluorescence analyses of tumor tissues following different treatment regimens. (a) Representative images of 4T1 tumor sections collected after various treatment: (i) H&E staining on day 16 post‐treatment, (ii) DCFH‐DA staining at 12 h post‐administration, (iii) TUNEL staining for apoptotic cells on day 16 post‐treatment, (iv) immunofluorescence staining for Hsp70 expression on day 16 post‐treatment, and (v) immunofluorescence staining for Hsp90 expression on day 16 post‐treatment. (b) Quantitative analysis of mean fluorescence intensity (MFI) for (i) ∙OH generation via DCFH‐DA staining, (ii) TUNEL signal, (iii) Hsp70 expression, and (iv) Hsp90 expression in tumor tissues after various treatments. Data are derived from the staining results shown in panels (a, ii)—(a, v) (n = 5 per group). Data are presented as mean ± standard deviation (n = 5).

3. Conclusion

In this study, we developed a plasmonically reinforced self‐sufficient nanozyme, designed as the plasmonic Pt‐CuO₂ nanozyme, for plasmon‐enhanced catalytic cancer therapy. The nanozyme consists of Pt NPs and CuO₂ nanodots co‐deposited on plasmonic AuSts, encapsulated within PEGylated ZIF‐8 MOFs. The prominent features of this nanozyme lie in its multi‐functional integration, which encompasses Pt/CuO₂‐based catalytic ∙OH generation, GSH depletion, self‐sufficient H₂O₂ supply, HSP downregulation‐enhanced mild PHT, plasmon‐enhanced catalytic activity, and TME responsive degradation. Following tumor accumulation, the acidic TME triggers degradation of the nanozymes, resulting in the release of Pt NPs and CuO₂ nanodots, along with in situ generation of H₂O₂ and Cu²⁺. The released Cu²⁺ depletes overexpressed GSH through direct oxidation, yielding Cu⁺, which together with Pt NPs catalyzes the production of substantial ∙OH from self‐supplied H₂O₂. Plasmonic heating and “hot electron” injection further enhance the ∙OH generation. Under 808 nm laser irradiation, the susceptibility of mitochondria to ROS leads to suppressed ATP biosynthesis and downregulation of HSP expression, thereby increasing thermal sensitivity and improving mild PHT efficacy. In vivo studies using a TNBC murine model demonstrated exceptional anticancer performance, attributable to the synergistic effect of enhanced ∙OH generation potentiated by GSH depletion, self‐supplied H₂O₂ and plasmonic enhancement. Our nanoyzme is fundamentally distinct from previously reported systems in its design, underlying mechanism, and therapeutic strategy [48]. This work represents a significant conceptual and methodological advancement, demonstrating promising potential for translational anticancer applications.

4. Experimental Section

4.1. Chemicals and Reagents

Zinc nitrate hexahydrate (Zn(NO₃)₂·6H₂O, 99%), 2‐methylimidazole (2‐mIM, 98%), copper chloride dihydrate (CuCl₂·2H₂O, 99.99%), and polyvinylpyrrolidone (PVP, (C₆H₉NO)_n, average molecular weight ≈ 10 kg/mol) were obtained from Tokyo Chemical Industry Co., Ltd. Chloroauric acid (HAuCl₄·4H₂O, 99% trace metals basis) was obtained from Shanghai Civi Chemical Technology Co. Ltd. 3,3′,5,5′‐Tetramethylbenzidine (TMB, 98%), rhodamine 6G (Rh6G, 99%), 5,5‐dimethyl‐1‐pyrroline N‐oxide (DMPO, 97%), and 5,5ʹ‐ dithiobis (2‐nitrobenzoic acid) (DTNB, 99%) were purchased from Aladdin Chemistry Co., Ltd. N, N‐dimethylformamide (DMF, anhydrous 99.8%), L‐ascorbic acid (AA, 98.0%) and hydrogen peroxide (H₂O₂, 30.0% (w/w)) were obtained from Sinopharm Chemical Reagent Co., Ltd. 1,2‐Distearoyl‐sn‐glycero‐3‐phosphoethanolamine‐N‐(methoxy(polyethyleneglycol)‐2000) (DSPE‐mPEG, average molecular weight ≈ 2 kg/mol, 95%) was purchased from Hunan Hua Teng Pharmaceutical Co., Ltd. Fetal bovine serum (FBS) and Roswell Park Memorial Institute 1640 medium (RPMI‐1640) were purchased from Biological Industries Israel Beit Haemek Ltd. JC‐1 mitochondrial membrane potential assay kit, 3‐(4,5‐dimethylthiazol‐2‐yl)‐2,5‐diphenyltetrazolium bromide (MTT) cell viability proliferation assay kit, 2,7‐dichloro‐fluorescin diacetate (DCFH‐DA) assay kit, Hoechst 33342, ATP assay kit, and Calcein‐AM/PI cell viability/cytotoxicity assay kit were purchased from Beyotime Biotechnology Co. Ltd. Ultrapure water (resistivity: 18.2 MΩ cm) used in all experiments was produced with a Millipore Direct‐Q3 UV system. Other reagents and solvents were of analytical grade and were used without any further purification, unless otherwise noted.

4.2. Characterization

Transmission electron microscopy (TEM) images were taken with an FEI Tecnai G² F20 S‐TWIN TMP transmission electron microscope at 200 kV. High‐angle annular dark‐field scanning transmission electron microscopy (HAADF‐STEM) imaging and energy‐dispersive X‐ray spectroscopy (EDS) analysis were performed on a Thermo Fisher Scientific Talos F200X scanning/transmission electron microscope at 200 kV. Scanning electron microscopy (SEM) images were acquired on a Hitachi Regulus 8230 field‐emission scanning electron microscope at 15 kV. Powder X‐ray diffraction patterns were recorded on a Rigaku D/Max 2500VB 18 kW X‐ray diffractometer at 40 kV and 250 mA using Cu Kα radiation (λ = 1.5406 Å). Optical extinction spectra were recorded on an Agilent Cary 5000 UV–Vis–NIR spectrophotometer. Hydrodynamic size and ζ‐potential measurements were performed using a Malvern Zetasizer Nano ZSE analyzer (ZEN3700). Electron spin resonance (ESR) spectra were obtained using a JES‐FA200 electron paramagnetic resonance spectrometer. Fourier Transform Infrared (FTIR) spectra were recorded using a Thermo Scientific Nicolet iS20 FTIR spectrometer. The concentrations of Au, Pt, and Cu in solutions and tissues were measured by an Agilent Cary 5100 inductively coupled plasma‐optical emission spectrometer (ICP‐OES) or a PerkinElmer NEXION 2000 inductively coupled plasma‐mass spectrometer (ICP‐MS). The cell viability measurements were conducted using MTT assay on a Tecan Spark multimode microplate reader. Cellular fluorescence images were taken on a Zeiss Axio Vert. A1 inverted fluorescence microscope.

4.3. Synthesis of Au Nanostars

Au nanostars (AuSts) were synthesized using a seed‐mediated growth method reported in our previous work [41, 42]. First, Au seeds were prepared following a well‐established protocol. Then, 1.5 g PVP was added to 15 mL DMF and ultrasonically dissolved, followed by sequential additions of 43 µL HAuCl₄·4H₂O (50 mm) and 20 µL pre‐prepared Au seeds. The reaction proceeded at room temperature for 3 h, and the reaction solution became dark blue, indicating the formation of AuSts. Finally, AuSts were purified by centrifugation and washing with water, which were stored in water at a concentration of 1.5 mg for following uses.

4.4. Synthesis of CuO₂ Nanodots

CuO₂ nanodots were synthesized according to a previously reported method [43]. First, CuCl₂·2H₂O (10 mm) was dissolved in 5 mL aqueous solution of 0.1 g/mL PVP, followed by sequential addition of 5.0 mL NaOH aqueous solution (20 mm) and 100 µL H₂O₂ (30.0% (w/w)). After 30 min of reaction, CuO₂ nanodots were purified by ultrafiltration and washed three times with water.

4.5. Synthesis of the Plasmonic Pt‐CuO₂ Nanozyme

First, Pt‐AuSts were synthesized through a wet‐chemistry reduction method. In brief, 2.5 µL of 48 mM H₂PtCl₆∙6H₂O was added to 4.0 mL AuSt aqueous suspension (500 µg/mL) under magnetic stirring, followed by 200 µL 100 mm AA introduced to initiate the reduction reaction. After thoroughly mixing, the reaction proceeded at 37 °C for 14 h, and the reaction mixture was centrifuged and washed with water, yielding Pt‐AuSts stored in water at 1.2 mg/mL. Afterward, 1.5 mL of 1.2 mg/mL Pt‐AuSts and 1.5 µL of 1.0 mg/mL CuO₂ nanodots were added to 1.0 mL methanolic solution of 1.32 m 2‐mIm, along with the addition of 2.0 mL 0.5 m Zn(NO₃)₂∙6H₂O. After 12 h of reaction at room temperature under magnetic stirring, the solution was centrifuged and washed three times with ethanol, yielding Pt‐CuO₂@AuSt@MOFs (denoted as plasmonic Pt‐CuO₂@MOFs) dispersed in ethanol at 1.0 mg/mL. Subsequently, 2.0 mg of DSPE‐mPEG was added to 10.0 mL ethanolic solution of plasmonic Pt‐CuO₂@MOFs obtained above. After sonication for 30 min, the reaction mixture was centrifuged and washed three times with ethanol, yielding PEGylated plasmonic Pt‐CuO₂@MOFs (denoted as plasmonic Pt‐CuO₂ nanozyme) that were dispersed in ethanol at 4.0 mg/mL for the following uses. Additionally, we also prepared PEGylated AuSt@MOFs (denoted as AuSt‐PEG) and PEGylated Pt‐AuSt@MOFs (denoted as plasmonic Pt nanozyme) using a similar synthetic protocol, but in the absence of Pt NPs/CuO₂ nanodots and CuO₂ nanodots, respectively.

4.6. Numerical Analysis of Plasmonic and Photothermal Properties

Numerical simulations of near‐field enhancement distribution and photothermal heating were carried out using the finite‐element method (FEM) within the COMSOL Multiphysics 5.6 software environment. The radio frequency (RF) model and heat transfer (HT) module were employed, following methodologies detailed in our previous work [22]. The AuSt was modeled as an Au nanosphere of a diameter of 53 nm containing ten spikes uniformly distribution on the 3D surface, and the Pt‐AuSt was constructed using the AuSt with each spike containing seven Pt semi‐spheres of a radius of 1.0 nm. The dimensional parameters and configuration of the AuSt and Pt‐AuSt were detained in Figure S3. For near‐field enhancement simulations, a 3D spherical domain with a radius of 350 nm was constructed. The target nanostructure (AuSt or Pt‐AuSt) was positioned at the origin (0, 0, 0). This was surrounded with a 250 nm‐thick water medium layer, followed by a 100 nm‐thick perfectly match layer (PML) to mitigate spurious reflection. For photothermal simulations, a cubic domain with a total edge length of 500 nm was used. The nanostructure (AuSt or Pt‐AuSt) was again placed at the origin. The outermost region comprised a 50 nm‐thick PML layer, with the space between the nanoparticle and the PML filled with water. A linearly polarized plane wave, polarized along the z‐axis and propagation along the x‐axis, served as the excitation source for near‐field analysis. For photothermal simulations, a continuous‐wave (CW) laser source was used. The dielectric function of Au was adopted from Johnson and Christy's experimental data [53, 54], while the dielectric constant of water was set to 1.77. The 3D simulation domain was discretized into tetrahedra elements using the built‐in algorithm integrated in the COMSOL software.

4.7. In Vitro Evaluation of the Photothermal Performance

To assess the photothermal performance of the plasmonic Pt‐CuO₂ nanozyme, various concentrations (0, 25, 50, 125, and 250 µg/mL) of the plasmonic Pt‐CuO₂ nanozyme was dispersed in ultrapure water in a microcentrifuge tube, which was irradiated with a 808 nm‐laser at a laser power density of 1.0 W/cm² for 10 min. The temperature was longitudinally monitored with an FLIR A35 infrared thermal camera. We also tested the photothermal effects of the aqueous suspension containing 250 µg/mL plasmonic Pt‐CuO₂ nanozyme at varied power densities (0.2, 0.4, 0.6, 0.8, and 1.0 W/cm²) under 808 nm laser. The photothermal stability of the plasmonic Pt‐CuO₂ nanozyme (250 µg/mL) was tested under 808 nm laser at a power density of 1.0 W/cm² for four consecutive laser on/off cycles. The photothermal conversion efficiency (η) was determined according to the previous reported protocol, described below: [55, 56]

$$\eta =\frac{hS\mathrm{\Delta }{T}_{\max }-Q_0}{I\left(1-{10}^{-A_{808}}\right)}$$ (1)

$$t=-\frac{m_s{C}_s}{hS}\ln \theta$$ (2)

$$\theta =\frac{T-T_{\mathrm{amb}}}{\mathrm{\Delta }{T}_{\max }}$$ (3)

where h is the thermal transfer coefficient, S is the surface area of the sample container, ∆T _max is the temperature difference between the steady‐state temperature (T _max) under 808 nm laser and the ambient temperature (T _amb), I is the laser power density, t is the cooling‐time following the laser switched off, m _s and C _s correspond to the suspension mass and thermal capacity (4.2 J/(g·°C)) of the aqueous solvent, respectively, and A ₈₀₈ is the absorbance at 808 nm of the suspension at the laser wavelength 808 nm. Q₀ is the heat dissipated by the solvent (water) absorbing the incident laser at 808 nm, which can be calculated by Q₀ = 5.4 × 10⁻⁴ × I (J/s).

4.8. Analysis of Electron Spin Resonance (ESR) Spectroscopy

ESR spectroscopy was utilized to detect ∙OH with DMPO as the trapping agent [49, 50]. The ESR spectra of DMPO/∙OH presented a characteristic quartet pattern with a peak intensity ratio of 1:2:2:1. The plasmonic Pt‐CuO₂ nanozyme was added at 40 µg/mL to 200 µL aqueous solution (pH 5.4) containing 1.0 µL DMPO in the presence or absence of 10 mm H₂O₂. For laser irradiation (+) groups, the samples were irradiated under 808 nm laser at 1.0 W/cm² for 5 min. After incubation for 0.5 h, 50 µL of the reaction mixture was taken to a capillary tube for ESR measurements.

4.9. In Vitro Analysis of OH Generation

The ∙OH generation ability of the plasmonic Pt‐CuO₂ nanozyme was assessed using a TMB colorimetric method under 808 nm laser or no laser irradiation, in which TMB was converted into oxidized TMB (oxTMB) with a characteristic absorption at 652 nm [11]. In brief, the plasmonic Pt‐CuO₂ nanozyme was added at varied concentrations (0, 10, 20, and 40 µg/mL) to 300 µL PBS (pH 5.4) containing 10 mm H₂O₂ and 10 mm TMB. After incubation for 5 min at room temperature, the absorbance at 652 nm of oxTMB was tested for ∙OH analysis. In addition, the ∙OH generation was examined at varied pH values (pH 5.4, 6.5, and 7.4) with other parameters kept identical. For studies on plasmon‐enhanced ∙OH generation, 5.0 µL of 1.0 mg/mL Pt‐AuSts were added to 200 µL PBS (pH 5.4) containing 10 mm TMB and 10 mm H₂O₂, which were kept under no laser or 808 nm laser (1.0 W/cm²) or under 808 nm laser with ice cooling for 5 min. The ∙OH generation was analyzed with TMB as the molecular probe as described above.

4.10. Analysis of the Catalytic Reaction Kinetics

Similarly, TMB was used as the molecular probe to analyze the catalytic reaction kinetics for ∙OH generation. First, to 2.0 mL PBS (pH 5.4) was added with 40 µL 10 mm TMB and 80 µL H₂O₂ of various concentrations (4, 8, 16, 32, and 64 mm), followed by the addition of 5.0 µL 1.0 mg/mL plasmonic Pt‐CuO₂ nanozyme. The optical absorption spectra of the reaction solution were longitudinally monitored every 2 s over a 10 min period, and the absorbance at 652 nm was used for the calculation of the concentration of the substrate H₂O₂. For quantitative analysis of the catalytic reaction kinetics, reaction rates (v₀ ) under various concentrations of H₂O₂ were calculated from the initial slope of TMB oxidation with time. Michaelis–Menten constant (K _m) and maximal reaction velocity (V _max) were determined through fitting the data to the Michaelis–Menten curve: [5]

$$v_0=\frac{V_{max}·\left[S\right]}{K_m+\left[S\right]}$$ (4)

$$\frac{1}{{\nu }_0}=\frac{K_m}{V_{max}}·\frac{1}{\left[S\right]}+\frac{1}{V_{max}}$$ (5)

where v ₀ is the initial reaction rate, and [S] is the concentration of the H₂O₂ substrate.

4.11. In Vitro Evaluation of the GSH Depletion Ability

The GSH depletion was tested with Ellman's reagent—DTNB as the molecular probe that reacts with GSH to yield yellow 2‐nitro‐5‐thiobenzoate acid with a characteristic absorption at 412 nm [50]. In brief, 1.0 mL of 10 mm GSH was added to 1.0 mL PBS (pH 5.4), followed by the addition of 0.5 mL 1.0 mg/mL plasmonic Pt‐CuO₂ nanozymes. The mixture was incubated at 37 °C, 50 µL of which was taken every 30 min and added with 0.2 mL 0.5 mm DTNB. The absorbance at 412 nm of the mixture was tested for quantitative analysis of GSH depletion. In addition, the GSH depletion was examined at varied pH values (pH 5.4, 6.5, and 7.4) with other conditions kept identical.

4.12. In Vitro Analysis of H₂O₂ Generation

The ability of the Pt‐CuO₂ nanozyme to generate H₂O₂ was assessed using KMnO₄ as the probe that reacts with H₂O₂ to generate Mn²⁺, along with the purple color changing to colorless [51]. Specifically, 1.0 mL aqueous solution of 0.1 m H₂SO₄ was added with 1.0 mL 50 µg/mL KMnO₄, followed by the addition of various concentrations (0.2, 0.4, 0.6, 0.8, and 1.0 mg/mL) of plasmonic Pt‐CuO₂ nanozymes. After incubation for 15 min, the absorption spectra were recorded from 400 to 650 nm.

4.13. Cell Culture and Animal Tumor Model

4T1 triple‐negative breast cancer cell line and L‐02 human normal liver cell line were obtained from the American Type Culture Collection (ATCC). The cells were cultured in RPMI‐1640 supplemented with 10% FBS and 1% penicillin‐streptomycin in a humidified incubator containing 5% CO₂ at 37 °C. All animal experiments were carried out in compliance with the requirements of laboratory animal welfare and ethics, and were approved by the Institutional Animal Care and Use Committee of the Central South University (Approval No. CSU‐2022‐0472). Female Balb/c nude mice (4 weeks old, 13–16 g) were purchased from Hunan SJA Laboratory Animal Co. Ltd. The 4T1 breast cancer xenograft mouse model was established by subcutaneously injecting 100 µL PBS containing 1.0 × 10⁶ 4T1 cells into the right hind limb of the mouse. When the 4T1 tumor volume reached approximately 120 mm³, the 4T1 breast tumor xenograft model was established and used for in vivo experiments.

4.14. Cytotoxicity Assay and In Vitro Anticancer Effect Evaluation

First, the biocompatibility of the plasmonic Pt‐CuO₂ nanozyme was assessed against 4T1 breast cancer cells and L‐02 normal liver cells by MTT assay. Specifically, cells were seeded into 96‐well plates and incubated at 1.0 × 10⁴ cells/well and incubated at 37 °C overnight for adhesion. Then, the cells were washed with PBS and incubated in fresh culture medium containing various concentrations of the plasmonic Pt‐CuO₂ nanozyme (0, 15, 30, 60, 120, and 180 µg/mL) for 24 h. For laser irradiation (+) groups, the treated cells were exposed to a 808 nm laser at a power density of 1.0 W/cm² for 5 min and cultured for another 4 h. Afterward, the cell culture medium was removed, and the cells were treated with 100 µL RPMI‐1640 containing 10 µL MTT reagent for 4 h, followed by removal of the culture medium removal and subsequent addition of 100 µL dimethyl sulfoxide. After shaking for 10 min, the absorbance at 570 nm of each well was measured for quantitative analysis of cell viability.

Further, the treated cells were also analyzed by a live/dead cell staining assay. Specifically, 4T1 cells were seeded into 96‐well plates at a cell density of 0.5 × 10⁴ cells/well, and cultured for 24 h for adhesion. The culture medium was replaced with fresh culture medium containing PBS or 180 µg/mL of AuSt‐PEG, plasmonic Pt nanozyme, and plasmonic Pt‐CuO₂ nanozyme. After 20 h incubation, the cells were irradiated with 808 nm laser at 1.0 W/cm² for min and subsequently cultured for 4 h for (+) groups. Afterward, the cells were washed with PBS and co‐stained with Calcien‐AM/PI, followed by fluorescence imaging.

For analysis of cellular uptake, 4T1 cells were seeded into 96‐well plates and cultured for 24 h for adhesion. Then, the culture medium was replaced with fresh RPMI‐1640 containing 30 µg/mL plasmonic Pt‐CuO₂ nanozymes fluorescently labeled with rhodamine 6G (plasmonic Pt‐CuO₂ nanozyme^Rh6G). After incubation for 0, 3, 6, or 9 h, the culture medium was removed, and the cells were washed twice with PBS. Subsequently, the cells were stained with Hoechst 33342 (nucleus, blue) for 10 min, followed by washing three times with PBS and subsequently examination by fluorescence imaging.

4.15. Analysis of the Mitochondria Membrane Potential

The mitochondrial membrane potential was examined using the JC‐1 mitochondrial membrane potential assay kit following the manufacturer's instruction [30]. 4T1 cells were seeded into 96‐well plates at a cell density of 0.5 × 10⁴ cells/well and cultured for 24 h for adhesion. Then, the culture medium was replaced with fresh RPMI‐1640 containing PBS, AuSt‐PEG, plasmonic Pt nanozyme, and plasmonic Pt‐CuO₂ nanozymes (60 µg/mL) for 12 h. For the (+) groups, the treated cells were irradiated with 808 nm laser at 1.0 W/cm² for 5 min. Afterward, the cells were washed twice with PBS and incubated in JC‐1 working solution for 20 min, which were observed by fluorescence microscopy.

4.16. Analysis of Intracellular GSH Depletion

Specifically, 4T1 cells were seeded into sixwell plates and cultured for 24 h for adhesion. The cells were incubated with the plasmonic Pt‐CuO₂ nanozyme at varied concentrations (0, 15, 30, 60, 120, and 180 µg/mL) for 6 h. Afterward, the culture medium was removed, and the cells were washed twice with PBS. Subsequently, 80 µL of 0.4% Triton‐X‐100 lysis buffer was added to lyse the cells, and after the lysates were centrifuged at 7500 rpm for 10 min, 50 µL of the supernatant was mixed with 200 µL0.5 mm DTNB. The absorbance at 412 nm was measured for quantitative analysis of GSH depletion.

4.17. Detection of Intracellular ·OH

Intracellular ∙OH generation was assessed using DCFH‐DA as the fluorescent probe. In brief, 4T1 cells were seeded into 96‐well plates and cultured for 24 h for adhesion. Then, the culture medium was replaced with fresh culture medium containing PBS, AuSt‐PEG, plasmonic Pt nanozyme, and the plasmonic Pt‐CuO₂ nanozyme at 60 µg/mL, and cultured for 6 h. For the (+) groups, the cells were irradiated under 808 nm laser at a power density of 1.0 W/cm² for 5 min. Then, the treated cells were added with 100 µL culture medium containing 10 µm DCFH‐DA and incubated for 30 min. Afterward, the culture medium was removed, and the cells were washed twice with PBS, which were examined by fluorescence microscopy.

4.18. Intracellular ATP Measurements

4T1 cells were seeded into sixwell plates, and cultured for 24 for adhesion. Then, the cells were incubated in RPMI‐1640 containing various concentrations of the plasmonic Pt‐CuO₂ nanozyme (0 (PBS), 15, 30, 60, 120, and 180 µg/mL) for 6 h. Afterward, the culture medium was removed, and the cells were washed twice with PBS, followed by the ATP measurement using the ATP assay kit according to the manufacturer's instruction.

4.19. Western Blot Analysis

Western blot analysis was performed to determine the expression levels of Hsp70 and Hsp90 in 4T1 cells following different treatments. Briefly, 4T1 cells were treated with PBS, AuSt‐PEG, plasmonic Pt nanozyme, and plasmonic Pt‐CuO₂ nanozymes at a concentration of 60 µg/mL for 24 h under 808 nm laser (1.0 W/cm², 5 min). After incubation, the cells were collected, washed, and lysed to extract total proteins. The protein concentrations were determined using a BCA assay to ensure equal loading. Subsequently, equal amounts of protein samples were separated by SDS‐PAGE and transferred onto a PVDF membrane. The membrane was then blocked with 5% non‐fat milk to prevent non‐specific binding. After blocking, the membrane was incubated with primary antibodies against Hsp70 and Hsp90, followed by incubation with an HRP‐conjugated secondary antibody. The protein bands were visualized using an enhanced chemiluminescence detection system. To ensure accuracy and reproducibility, β‐actin was used as an internal control for normalizing protein expression levels.

4.20. Hemolysis Experiment

Red blood cells (RBCs) were collected from fresh mouse blood, and washed five times with PBS. Then, different concentrations of plasmonic Pt‐CuO₂ nanozymes (12.5, 25, 50, 100, 200, and 400 µg/mL) were added into RBC suspensions. PBS and ultrapure water were used as negative control and positive control, respectively. After incubation at 37 °C for 24 h, the samples were centrifuged at 10 000 rpm for 10 min, and the supernatant was collected for the absorbance measurement. The hemolysis rate was calculated by the following formula: [52]

$$\mathrm{Hemolysis}\mathrm{rate}\left(\%\right)=\frac{A_{\mathrm{s}}-A_{\mathrm{n}-\mathrm{ctr}}}{A_{\mathrm{p}-\mathrm{ctr}}-A_{\mathrm{n}-\mathrm{ctr}}}\times 100$$ (6)

where A _s, A _n‐ctr, A _p‐ctr are the absorbance at 416 nm of the samples, negative control, and positive control, respectively.

4.21. In Vivo Photothermal Measurements

For in vivo photothermal experiments, 200 µL of PBS or 1.0 mg/mL plasmonic Pt‐CuO₂ nanozyme in PBS was intravenously administered into the 4T1 tumor‐bearing mice. The mice were then exposed to 808 nm laser at 1.0 W/cm² for 5 min at 12 h post‐administration. The temperature profile and infrared thermal images of the mouse tumors were longitudinally recorded with an FLIR A35 infrared thermal camera over the 5 min‐laser irradiation period.

4.22. Evaluation of In Vivo Therapeutic Efficacy in TNBC Breast Tumor Model

Once the tumor size reached 120 cm³, the 4T1 tumor‐bearing mice were randomly divided into four groups (n = 5 per group): (1) PBS (200 µL) + laser, (2) AuSt‐PEG (20 mg/kg) + laser, (3) plasmonic Pt‐CuO₂ nanozyme (20 mg/kg), and (4) plasmonic Pt‐CuO₂ nanozyme (20 mg/kg) + laser. For the laser irradiation (+) groups, the mice were irradiated with an 808 nm laser at 1.0 W/cm² for 5 min at 12 h post‐administration. The tumor volume and body weight of the mice were measured every other day over a 16‐day period. The tumor volume was calculated using the following formula: V = L × W²/2, where L and W are the length and width of the tumors, respectively. Relative tumor volume was calculated as V/V₀, where V₀ is the initial tumor volume. The in vivo ∙OH generation in tumor tissues was assessed by DCFH‐DA staining at 12 h post‐administration. On day 16, the mice were sacrificed, and the tumor and major organs (heart, liver, spleen, lung, kidney) were harvested for weighting, photographing, hematoxylin and eosin (H&E) staining, HSP70/HSP90 immunofluorescence staining, and terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining. Blood was collected from on day 16 after various treatments for blood biochemical analysis and liver/kidney function examination, including alanine transaminase (ALT), alkaline phosphatase (ALP), blood urea nitrogen (BUN), aspartate transferase (AST), and lactate dehydrogenase (LDH).

4.23. Statistical Analysis

All experiments were carried out with at least three repeats independently, and all results were presented as mean ± standard deviation (SD). Data analysis was performed using Origin 2019b and GraphPad Prism 8.3.0 Software. Statistical comparison between different groups was carried out using a one‐tail Student's t‐test. Statistical values are indicated as p values with p < 0.05 considered significantly different: ^* p <0.05, ^** p <0.01, ^*** p <0.001, and ^**** p <0.0001.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71394‐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.