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Remodeling tumor microenvironment using prodrug nMOFs for synergistic cancer therapy

Journal of Nanobiotechnology volume 23, Article number: 123 (2025) Cite this article

Abstract

Metal–organic frameworks (MOFs) hold tremendous potential in cancer therapy due to their remarkable structural and functional adaptability, enabling them to serve as nanocarriers for biopharmaceuticals and nanoreactors for organizing cascade bioreactions. Nevertheless, MOFs are predominantly utilized as biologically inactive carriers in most cases. Developing nanoscale prodrug MOFs suitable for biomedical applications remains a huge challenge. In this study, we have designed a novel prodrug nano-MOFs (nMOFs, named DCCMH) using metformin (Met) and α-cyano-4-hydroxycinnamic acid (CHCA) as ligands for coordination self-assembly with CuCl2, followed by loading of doxorubicin (DOX) and surface modification with hyaluronic acid (HA). Upon internalization by cancer cells, DCCMH releases Cu2+/+, CHCA, Met, and DOX in response to high levels of glutathione (GSH) and hydrogen peroxide (H2O2) within the tumor microenvironment (TME); Cu+ catalyzes the conversion of H2O2 to ·OH via the Fenton reaction while it was oxidized to Cu2+, which was subsequently further de-consumed of GSH; CHCA induces a further decrease in intracellular pH and promotes Fenton reactions by inhibiting lactate efflux; Met up-regulates tyrosine kinase activity and enhances the chemotherapy of DOX. With the ability to synergistically combine chemo/chemodynamic therapy (CT/CDT) and remodel the TME, the DCCMH NPs inhibit murine hepatoma effectively. This study presents a feasible strategy for fabricating prodrug nMOFs which are capable of remodeling TME to improve efficacy through synergistic cancer therapy.

Graphical abstract

Introduction

The TME plays a pivotal role in fostering the proliferation and dissemination of malignant cells [1, 2]. It is characterized by elevated levels of H2O2, overexpressed GSH, moderate acidity, hypoxia, and vigorous metabolism [3, 4]. To effectively eradicate tumors, it is imperative to remodel the TME in conjunction with tumor treatment strategies [5,6,7]. Numerous methodologies have been documented for restructuring the TME, encompassing the enhancement of hypoxia conditions [8], regulation of the tumor extracellular matrix [9], utilization of tumor-associated fibroblast-targeting nanotherapy [10], and reconfiguration of the tumor vascular system [11]. However, solely remodeling the TME is not sufficient to eradicate cells, the incorporation of other therapeutic agents is also needed [12,13,14,15,16,17,18]. The field of nanotechnology has witnessed noteworthy advancements, particularly in the realm of nanomedicines with multimodal synergistic therapy [19,20,21,22,23,24]. A particularly auspicious approach in this domain is the utilization of nMOFs, which are hybrid materials renowned for their stable structures, controllable components, and efficient encapsulation of small-molecule drugs [25,26,27]. These distinctive properties have propelled nMOFs into the limelight of nanomedicine, garnering considerable attention and acclaim [28,29,30].

nMOFs, known for their self-assembled structures composed of metal ions and organic ligands [31], have exhibited immense potential in synergistically inhibiting tumor growth by combining the anticancer properties of metal ions with the delivery of chemotherapy and immunotherapy drugs [32,33,34]. However, the current nMOFs suffer from limited bioavailability, primarily attributed to the use of inactive and inflexible precursor ligands. To tackle this, researchers have recently proposed the incorporation of biologically active drug molecules as precursor ligands, giving rise to prodrug nMOFs [35]. These prodrug nMOFs selectively release their encapsulated drugs under specific stimuli, effectively preventing premature drug leakage and enhancing the overall anticancer efficacy of the components [36, 37]. Despite the promising nature of this approach, the applications of prodrug nMOFs in antitumor combination therapy remain under-reported. To enhance the efficacy of prodrug nMOFs, we advocate for their construction in conjunction with the remodeling of the TME. By specifically targeting and modifying the TME, we can create a more conducive environment for the release and action of the drugs encapsulated within the prodrug nMOFs. This innovative approach holds the potential to augment the therapeutic effects of nMOFs and elevate their overall antitumor activity.

Herein, we successfully synthesized GSH-responsive prodrug nMOFs through the coordination of CuCl2·2H2O with CHCA and Met under solvothermal conditions. The chemotherapeutic agent, DOX, was encapsulated into the nanoporous structure, and the HA was surface-functionalized onto the nMOFs to improve biocompatibility and tumor-specific targeting. As depicted in Scheme 1: (1) The as-prepared HA-coated and DOX-loaded coordination nMOFs were termed DCCMH, which were internalized by cancer cells via the specific interaction between HA and receptors on the cell membrane. (2) Subsequently, the DCCMH underwent gradual degradation within cancer cells by high concentrations of GSH, leading to the liberation of DOX, CHCA, Met, and Cu2+/+. (3) Cu+ catalyzes the conversion of H2O2 to ·OH via the Fenton-like reaction while it was oxidized to Cu2+, which was subsequently further de-consumed of GSH. This catalytic process facilitated the conversion of intracellular H2O2 into highly toxic ·OH. (4) The monocarboxylic acid transporter (MCT) inhibitor, CHCA, effectively impeded the efflux of lactic acid, thereby inducing an elevation in intracellular acidity. Consequently, this enhanced the efficiency of the Fenton-like reaction, intensifying oxidative damage to tumor cells. (5) Met triggered the up-regulation of AMP-activated protein kinase (AMPK), causing disruption to normal metabolic pathways and augmenting the chemotherapeutic sensitivity of tumor cells to DOX. The synergistic effect resulting from the depletion of GSH, regulation of acidity/metabolism, and the reinforcement of CT/CDT induced substantial mitochondrial damage and disturbed the redox homeostasis of cancer cells, ultimately exerting robust inhibitory effects on tumors.

Scheme 1.
scheme 1

Schematic illustration of a the fabrication procedure of the HA-coated and DOX-loaded coordination nMOFs, and b the underlying mechanism of DCCMH for synergistic cancer therapy

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Results and discussion

Characterizations of DCCMH

Firstly, Cu-based nMOFs (CCM) were synthesized by a solvothermal method using CuCl2·2H2O, CHCA, and Met. Subsequently, DOX was loaded into CCM to obtain DCCM. Finally, HA was modified onto the surface of DCCM through electrostatic interactions, forming DCCMH. Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) images revealed that CCM and DCCMH NPs possessed a distinctive porous snowflake-like morphology, measuring approximately 225 ± 10 nm in diameter, with a spatial lattice dimension of 0.5 nm for CCM (Fig. 1a, b). The similar morphology of DCCM and CCMH (HA was modified onto the surface of CCM) was observed (Figure S1). Elemental analysis through energy-dispersive X-ray spectroscopy and element mapping confirmed the presence of C, N, O, and Cu elements within the CCM NPs (Fig. 1c). Dynamic light scattering analysis revealed the hydrodynamic diameters of 190.6 nm, 223.2 nm, and 248.4 nm, with polydispersity indices (PDI) of 0.085, 0.123, and 0.118 for CCM, DCCM, and DCCMH NPs, respectively (Fig. 1d). Furthermore, the stability test indicated that DCCMH NPs exhibited excellent stability in various physiological solutions, with the particle size remaining essentially unaltered in the absence of acidic pH and GSH stimulation (Figure S2). Notably, the sizes of the modified nanoparticles exhibited a gradual increase, accompanied by a corresponding decrease in zeta potential (Fig. 1e), due to the loading of DOX and surface modification of HA. The Fourier-transform infrared (FTIR) spectra revealed the coordination of Cu ions with Met and CHCA in CCM, and the blue shift characteristic peak proved that Cu ions coordinated with the guanidine group in Met and the carboxyl group in CHCA, ultimately forming a metal–organic complex with porous structure. Specifically, the peaks at 1300 cm−1 (–C–N), 1582 cm−1 (–COO), 1644 cm−1 (C=NH), 2208 cm−1 (–C≡N), 3300 cm−1 (Ph–OH), and 3450 cm−1 (-NH2) were indicative of these coordination interactions (Fig. 1f). The X-ray diffraction (XRD) pattern unequivocally demonstrated the crystalline nature of CCM, exhibiting a spatial configuration similar to CuO and CuN crystals and a characteristic XRD peak at a 2θ angle of 8° (Figs. 1g and S3), which combined with the high-resolution transmission (HR-TEM) images demonstrated the MOF structure. Furthermore, we determined the specific surface area of CCM and DCCM using the Brunauer–Emmett–Teller (BET) method (Figure S4). CCM showed a surface area of 46.28 m2 g−1 with a pore volume of 1.5 cm3 g−1, which decreased to 26.39 m2 g−1 with a pore volume of 0.052 cm3 g−1 after loading with DOX, further suggesting the characteristically porous MOF structure of the as-prepared CCM. Owing to the porous crystalline bulk structure, the calculated loading and encapsulation efficiencies of DOX in CCM nMOFs were determined to be 8.12% and 48.63%, respectively. The FTIR spectra displayed in Figure S5 confirmed the successful HA-functionalization and DOX-loading. The characteristic UV–Vis absorption band of DOX was observed in DCCMH (Fig. 1h). Moreover, to elucidate the element valence states, as shown in Fig. 1i, the X-ray photoelectron spectroscopy (XPS) analysis implied the characteristic signals of Cu, C, O, and N within the DCCMH. It is worth noting that Cu mainly exists in the form of Cu (I) in DCCMH, with a small amount of Cu (II). The presence of satellites around 942.0 and 961.2 eV is possibly due to the p → d hybridization of unsaturated d orbitals, indicating the presence of Cu (II) centers. The second set at 932.5 and 952.5 eV comes from Cu (I) centers, which usually have lower Cu2p3/2 and Cu2p1/2 binding energy due to the lower oxidation state [38]. In addition, the presence of satellites near 647.5 and 719.2 eV may represent the Auger peak corresponding to Cu.

Fig. 1
figure 1

a SEM images and b TEM images of CCM and DCCMH NPs (insert: the HR-TEM image of CCM). c Elemental mapping of DCCMH NPs. d The distribution of hydrodynamic diameters and e zeta potential of CCM, DCCM, CCHM, and DCCMH. (f) FTIR spectra of Met, CHCA, and CCM. g The XRD pattern of CCM NPs. h The UV–Vis absorption spectra of DOX, CCMH, and DCCMH. i The XPS spectrum of CCM NPs (insert: the high-resolution spectra of the Cu2p orbitals)

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GSH depletion, ·OH generation, and cytotoxicity assessments

The preservation of an optimal redox equilibrium is paramount for the survival of cells, and any disruption to this delicate balance can effectively impede the growth and proliferation of cancer cells [4]. Consequently, we assessed the capability of DCCMH to deplete GSH and generate ·OH. Figure 2a demonstrates a gradual decline in GSH content with increasing concentrations of DCCMH. This reaction can primarily be ascribed to the reduction of Cu2+ within DCCMH to Cu+ by GSH (Figures S6a and S6b). The DCCMH NPs undergo complete degradation within 6 h when the GSH concentration is 10 mM and the DCCMH concentration is 80 μg mL−1 (Figure S6c). Subsequently, as DCCMH NPs degrade, the encapsulated drugs are rapidly released, resulting in the release of up to 78% of DOX (Fig. 2b). This underscores the superior GSH-dissipative and responsive drug-release behavior of DCCMH NPs, as well as their capacity to disrupt the redox equilibrium of the TME by depleting GSH. Furthermore, we analyzed the release of CHCA and Met by observing their characteristic UV–Vis absorption spectra. To prevent any interference from the absorption spectrum of DOX on the determination of CHCA and Met absorption spectra, we opted to observe the changes in the absorption spectra of CCM without the presence of DOX before and after the addition of GSH. Subsequently, it was observed that the distinct absorption peaks of CHCA and Met were present in the absorption spectra after the degradation of CCM NPs in the GSH solution, indicating the successful release of CHCA and Met from the degraded CCM (Figure S7). This observation further verifies that the structure and effectiveness of Met and CHCA remained intact during the high-temperature synthesis of CCM. Then, the ·OH produced by a Fenton-like reaction of H2O2 with Cu+ was measured using methylene blue (MB) as a probe [39]. As shown in Fig. 2c, a more significant decrease of the MB absorbance in the MB + H2O2 + DCCMH + GSH group than in other groups indicates the remarkable CDT effect of DCCMH under the condition when both GSH and H2O2 are present. To effectively eliminate cancer cells, it is crucial to maintain continuous production of ·OH, as they possess a short half-life and easily lose activity [40]. Consequently, we investigated the ·OH production over time. As illustrated in Fig. 2d, in the presence of H2O2 and GSH, the absorbance of MB gradually decreased, signifying that ·OH was continuously and gradually generated in this system, ensuring its sustained cytotoxicity. Electron spin resonance (ESR) spectroscopy further validated the efficient production of ·OH, using DMPO as a trapping agent (Fig. 2e). The results showed that the signal of ·OH (1:2:2:1) was very weak without the addition of GSH, but its signal was significantly enhanced with the addition of GSH, which was mainly due to the degradation of DCCMH under the action of GSH to release a large amount of Cu+. However, in the absence of degradation of DCCMH, the Cu+ exposed on the surface was insufficient to catalyze the generation of ·OH from H2O2. The above results lay the foundation for DCCMH as an antitumor nanomedicine, which achieves antitumor effects by disrupting redox homeostasis to amplify oxidative stress.

Fig. 2
figure 2

a The relative GSH content treated with different concentrations of DCCMH for 6 h. b The DOX release profile of DCCMH under different conditions. c The UV–Vis absorption spectra of MB in different solutions. d The degradation of MB at different time intervals mediated by DCCMH. e The ESR spectra of H2O2, H2O2 + DCCMH, and H2O2 + DCCMH + GSH using DMPO as the trapping agent. f-i Cell viability assay of HL7702 cells and HepG2 cells incubated with different samples at varying concentrations for f 24 h of HL7702 cells, g 48 h of HL7702 cells, h 24 h of HepG2 cells, i 48 h of HepG2 cells (n = 6). (*p < 0.05, **p < 0.01, ***p < 0.001)

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Subsequently, the MTT assay was conducted to appraise the cytotoxicity of DCCMH towards cancer cells. As delineated in Fig. 2f, g, diverging from DOX, DCCMH exhibited negligible toxicity towards the human normal hepatic (HL7702) cells, whilst manifesting considerable cytotoxicity towards human hepatocellular carcinoma (HepG2) cells (Fig. 2h, i). DOX indiscriminately attacks various cells, whereas the specificity of DCCMH towards HepG2 cells, which overexpress the CD44 receptors on the surface, facilitates its specific uptake in HepG2 cells. Consequently, within the acidic environment of cancer cells and in the presence of high concentrations of GSH-H2O2, DCCMH rapidly degrades and releases synergistic anticancer drugs. The viability of HepG2 cells gradually declined with escalating drug concentration and incubation duration, wherein a mere 14.5% of HepG2 cells persevered following 24 h of incubation with 80 μg mL−1 of DCCMH (containing 6.4 μg mL−1 of DOX). Furthermore, the IC50 of DCCMH on HepG2 cells stood at 11.9 μg mL−1, underscoring its exceedingly potent capacity to annihilate hepatocellular carcinoma cells. Meanwhile, to more intensively assess the synergistic anticancer efficiency of DCCMH, the combination index (CI, CI < 1 indicates synergism, CI = 1 indicates additive effect, and CI > 1 indicates antagonism) was analyzed [41]. The CI value was determined by the formula: CI = DDOX/IC50(DOX) + DCCMH/IC50(CCMH), where IC50(DOX) and IC50(CCMH) represent the IC50 values under monotherapy (DOX or CCMH). DDOX and DCCMH are the concentrations of DOX and CCMH corresponding to the IC50 of DCCMH combination therapy. The results showed that the CI of DCCMH was 0.79, which strongly proved that DCCMH achieved good synergism of CDT and CT. Finally, DCCMH evinced commendable lethality against three additional cancer cell strains, namely Hela cells, U87MG cells, and 4T1 cells (Figure S8).

In vitro therapeutic efficacy

Next, we thoroughly elucidated the mechanism of action of DCCMH on HepG2 cells. Initially, we quantified the cellular internalization of DCCMH using advanced techniques such as flow cytometry and confocal laser scanning microscopy (CLSM) analysis. Remarkably, the results depicted in Figs. 3a and S9 demonstrated the profound translocation of DOX from DCCMH into the nucleus. Furthermore, the fluorescence signals emitted by DOX exhibited a steady augmentation within the HepG2 nucleus over time, indicative of a time-dependent elevation in the uptake of DCCMH NPs by HepG2 cells. Notably, the cell uptake of DCCMH reached maximal after 6 h, astonishingly presenting a ninefold increase in fluorescence signals compared to the control group (Figure S10). This enhanced cellular uptake of DCCMH NPs by HepG2 cells can predominantly be attributed to passive uptake based on the enhanced permeability and retention effect of nanomedicines as well as highly selective affinity-based active targeted uptake between HA in DCCMH and CD44 receptors abundantly expressed on the surface of HepG2 cells, which was proved by the CD44 blocking assay using HA (Figure S11) [42].

Fig. 3
figure 3

a CLSM images depicting the internalization of DCCMH by HepG2 cells at varying time intervals. b The intracellular total GSH levels under different groups of incubation (n = 6). c The intra/extracellular lactic acid content under different groups of incubation (n = 6). d CLSM images of HepG2 cells treated with different samples and stained with pH fluorescent probe (BCECF), Scale bar: 100 μm. e The intracellular ROS detection through DCFH-DA staining, Scale bar: 50 μm. f The mean fluorescence intensity values of intracellular DCFH (n = 3). g CLSM images of HepG2 cells treated with different samples and stained with JC-1, Scale bar: 100 μm. h Changes in the mitochondrial membrane potential of HepG2 cells (n = 6). i Analysis of AMPK enzyme activity in different treatment groups (n = 6). j Live/dead staining images of HepG2 cells after different treatments, Scale bar: 100 μm. k Flow cytometry analysis of HepG2 cell apoptosis. l Analysis of apoptosis time in different groups (Concentrations of drugs selected for cellular experiments: DOX: 4.8 μg mL−1, CCMH: 55.2 μg mL−1, and DCCMH: 60 μg mL.−1). (*p < 0.05, **p < 0.01, ***p < 0.001)

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Given the considerable degradation of DCCMH induced by GSH in solution, we proceeded to assess the depletion of intracellular GSH by DCCMH. As depicted in Fig. 3b, the levels of intracellular GSH decreased by 45% in the treatment groups of CCMH and DCCMH in comparison to the PBS group. Following the degradation of CCMH and DCCMH, their constituents CHCA, Met, and Cu+ were gradually liberated into the cancer cells. CHCA is specifically employed to inhibit the efflux of intracellular lactate [16, 17]. Hence, we scrutinized the intra/extracellular levels of lactate and the intracellular changes in acidity. As illustrated in Fig. 3c, the groups treated with CCMH and DCCMH exhibited significant alterations in the intra/extracellular content of lactic acid compared to the PBS and DOX groups. Notably, the DCCMH group manifested a remarkable twofold increase in intracellular lactic acid content, while simultaneously experiencing a substantial reduction of approximately 80% in extracellular lactic acid. The accumulation of intracellular lactic acid consequently led to a decline in intracellular pH, which was measured using the pH-sensitive fluorescent probe BCECF that exhibits green fluorescence positively correlated with pH level. As demonstrated in Fig. 3d, the CCMH and DCCMH groups displayed low fluorescence intensities of BCECF compared to the PBS and DOX groups. The Fenton reaction is more likely to occur under acidic conditions, therefore, the elevation of acidity in cancer cells is beneficial for the intracellular Fenton reaction to occur [20].

The underlying principle of CDT revolves around the Fenton-like reaction for the intracellular generation of substantial quantities of toxic ·OH, which effectively induces oxidative damage to the mitochondria, ultimately leading to the eradication of cancer cells [43]. In light of this, we assessed the intracellular ·OH levels and mitochondrial membrane potential changes. As illustrated in Figs. 3e and S12, the CCMH and DCCMH groups exhibited heightened DCFH fluorescence signals and generated significantly more ·OH compared to the PBS and DOX groups. Quantitative analysis of the average fluorescence intensity of DCFH revealed 73.68% in the DCCMH group, which was approximately 15-fold higher than that of the PBS group (Fig. 3f). By the efficient reduction-degradation process in cancer cells and the subsequent increase in acidity, the Cu+ ions released by DCCMH reacted efficiently with H2O2, continuously releasing reactive oxygen species (ROS) and facilitating a highly effective CDT effect. The changes in mitochondrial membrane potential, assessed using the membrane potential detection kit (JC-1), confirmed the damage inflicted upon cancer cells by CDT action (Figs. 3g and S13). Analysis of the fluorescence ratio of JC-1 dimers to JC-1 monomers (R/G) demonstrated a gradual decrease in the R/G ratio across the different incubation groups, as depicted in Fig. 3h. This decline in the R/G ratio signifies a reduction in mitochondrial membrane potential and an enhancement of oxidative stress in cancer cells. Furthermore, it has been discovered that Met can modulate the activity of tyrosine kinase in cancer cells, particularly in digestive carcinoma, thereby promoting cancer therapy [12,13,14]. As shown in Fig. 3i, compared with the PBS and DOX groups, both CCMH and DCCMH significantly up-regulated AMPK expression in HepG2 cells, thereby exerting a profound influence on cell metabolism and synergistically enhancing chemotherapy [15].

Subsequently, cell viability and apoptosis were evaluated using live/dead staining analysis and apoptosis assay. HepG2 cells were subjected to fluorescence imaging after staining with Calcein-AM (indicating live cells, depicted as green fluorescence) and propidium iodide (PI) (indicating dead cells, depicted as red fluorescence). Notably, HepG2 cells subjected to DCCMH exhibited significant cell apoptosis compared to the control groups (Figs. 3j and S14), showing a 67.1% rate of apoptosis, as quantitatively assessed by flow cytometry (Fig. 3k). This increase was predominantly observed in the initial stage of apoptosis, as indicated by the presence of cells in the Q3 region (Fig. 3l). Concurrently, the apoptosis cycle assay revealed that DCCMH could impede the progression of cancer cells in the G2/M phase (Figure S15). The amalgamation of TME-remodeling, Cu+-mediated CDT, and DOX-mediated chemotherapy synergistically contributed to the potentiation of apoptotic signaling, resulting in a heightened incidence of programmed cancer cell demise.

In vivo therapeutic efficacy

The biodistribution and clearance dynamics of DCCMH NPs within H22 tumor-bearing mice were meticulously examined through the strategic integration of Cy5.5 onto the DCCMH nanoplatforms. Notably, the fluorescence signals emitted by the DCCMH-Cy5.5 exhibited a gradual and substantial accumulation within the tumor milieu over an 8 h period, ultimately culminating in an extensive and enduring retention of up to 28 h (Figures S16a and S16b). Ex vivo fluorescence imaging of major organs further substantiated this observation, as evidenced by the conspicuously robust fluorescence signals discernible within the tumor regions post-intravenous administration of DCCMH-Cy5.5 after 28 h, while relatively strong fluorescent signals were also seen in kidneys tissues, which could be attributed to the eventual metabolism of the DCCMH by the kidneys after it has passed through the bloodstream in vivo (Figure S16c). Conversely, other tissues (heart, liver, spleen, lungs) have weaker fluorescence signals. These results demonstrated that DCCMH could accumulate effectively in tumor regions for a long time, which was conducive to its long-term therapy for cancers.

The in vivo therapeutic efficacy of the combination treatment was then assessed through the intravenous administration of DCCMH using H22 tumor-bearing mice. The H22 cell line is a widely utilized murine hepatocellular carcinoma cell line that overexpresses the CD44 receptor. It is suitable for establishing a fully immunocompetent hepatocellular carcinoma xenograft model [43,44,45,46]. Once the tumor size reached approximately 100 mm3, the H22 tumor-bearing mice were randomly divided into four groups (n = 7/group): (I) PBS (blank control), (II) DOX, (III) CCMH, (IV) DCCMH. The drugs were administered intravenously every 4 days (2 mg kg−1), and the mice were monitored for 18 consecutive days, with photographs taken, weights recorded, and tumor volumes measured every 3 days (Figs. 4a and S17-20). Monotherapy exhibited limited efficacy due to drug resistance and the rapid stress response of cancer. Remarkably, the DCCMH treatment effectively suppressed tumor growth, exhibiting an inhibition rate of 74.24%, surpassing other treatment modalities with statistically significant differences (Figs. 4b-e, S21, and S22). Extended survival time after various treatments was also observed, as shown by the survival curve in Figure S23. The mice died when the diameter of the tumor exceeded 15 mm according to NIH guidelines. The alterations in body weight exhibited no noteworthy aberrations across all experimental cohorts (Fig. 4f), signifying the absence of evident systemic toxicity. Furthermore, to more intensively assess the mechanisms of DCCMH, the activity of AMPK enzymes in isolated H22 tumor tissue was analyzed. As shown in Fig. 4g, compared with the PBS and DOX groups, both CCMH and DCCMH significantly up-regulated AMPK enzyme activity in H22 tumors, further promoting chemotherapeutic efficacy [47].

Fig. 4
figure 4

a Schematic illustration of the therapeutic intervention in the H22 subcutaneous tumor-bearing mouse model. b Respective tumor growth curves for each tumor-bearing mouse in different groups. c The average tumor growth patterns exhibited by H22 tumor-bearing mice following different treatments. d The weight of extracted H22 tumors after different treatments. e The photographs of extracted H22 tumors. f The changes in body weight of H22-tumor-bearing mice in different treatment groups. g Analysis of AMPK enzyme activity in H22 tumor tissue from different treatment groups (n = 5). h H&E staining, TUNEL labeling, and Ki-67 staining of tumors in mice subjected to various treatments for 18 days. All Error bars represent mean ± S.D. n = 5 biologically independent samples. (*p < 0.05, **p < 0.01, ***p < 0.001)

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Additionally, histopathological analyses of extracted tumor tissues were performed using hematoxylin & eosin (H&E), terminal deoxynucleotidyl transferase-mediated nick end labeling (TUNEL), and Ki67 staining to assess morphology, apoptosis, and proliferation of tumor cells (Fig. 4h). The H&E-stained tumor sections exhibited a significant decrease in cell density in the DCCMH treatment group. The intense green fluorescence observed in TUNEL staining and the weak red fluorescence observed in Ki67 staining suggested that DCCMH-mediated synergistic therapy induced significant apoptosis and inhibition of tumor cell proliferation. The main organs including the heart, liver, spleen, lungs, and kidneys were subjected to histological examination through H&E staining after all interventions, and the results indicated no significant organ toxicity within all experimental groups (Figure S24).

Finally, the biosafety of DCCMH was assessed. The hemolysis test (Figure S25) revealed that DCCMH exhibited negligible hemolytic activity on red blood cells across various concentrations (5–160 μg mL−1). Remarkably, even at a high concentration of 160 μg mL−1, the corresponding hemolysis rate was merely approximately 3%, underscoring the excellent biosafety profile of DCCMH. Subsequently, blood samples of mice with different treatments were collected for biochemical analysis and hematological evaluations. Notably, there were no significant differences between the experimental and the control groups in terms of liver function, renal function, or hematological parameters (Figures S26 and S27).

Conclusion

In summary, we have devised an innovative nMOF (DCCMH) utilizing prodrugs, aimed at effectively regulating the tumor microenvironment. Upon exposure to elevated GSH levels, DCCMH gradually breaks down, releasing Cu2+/+, CHCA, Met, and DOX into cancer cells, instigating a cascade of antitumor responses: (1) Cu+ catalyzes the transformation of H2O2 to ·OH through the Fenton reaction as it oxidizes to Cu2+, subsequently depleting GSH, leading to oxidative damage in cancer cells; (2) CHCA hinders lactic acid exocytosis, elevating intracellular acidity and intensifying the Fenton-like reaction; (3) Met triggers the AMPK pathway, diminishing tumor cell resistance to DOX. These characteristics promote collaborative TME restructuring and achieve remarkable antitumor efficacy, resulting in a tumor suppression rate of 74.24%. This study presents a fresh approach for the uncomplicated synthesis of prodrug MOFs capable of initiating a sequence of responses within cancer cells for TME regulation. Furthermore, this approach establishes a theoretical foundation for the clinical implementation of combination tumor therapy.

Experimental section

Materials and chemicals

All reagents used in this study were of analytical grade and were obtained from reputable suppliers. α-cyano-4-hydroxycinnamic acid, CuCl2·2H2O, glutathione, hydrogen peroxide (30%), and methylene blue were bought from Aladdin-Reagent Co. Ltd. (China). Metformin and hyaluronic acid (Mw < 10 kDa) were obtained from Coolaber Science & Technology Co. (Beijing, P. R. China). Doxorubicin was bought from HEOWNS-Reagent Co. Ltd. (Tianjin, P. R. China). Dimethyl sulfoxide was bought from Chengdu Chron Chemical Co. Ltd. Hoechst 33258, reduced GSH assay kit, Annexin V-FITC/PI kit, mitochondrial membrane potential kit (JC-10 Assay), 2′,7′-Dichlorofluorescin diacetate, and fetal bovine serum were purchased from Solarbio Science & Technology Co., Ltd. The Lactic acid content kit was purchased from Nanjing Jiancheng Biotechnology Co., Ltd. (Nanjing, P. R. China).

Preparation of CCM NPs

CuCl2·2H2O (0.1 mM), CHCA (0.02 mM), and Met (0.08 mM) were mixed and dissolved in 4 mL solution (VH2O: Vethanol = 1:1) with 30 μL of triethylamine, followed by ultrasonication to fully dissolve and transferred to a 15 mL Teflon-lined autoclave and heated at 100 ℃ for 3 h. After the reaction, the samples (CCM NPs) were washed with ultrapure water and methanol to remove impurities.

Preparation of DCCM and DCCMH NPs

Aqueous DOX (1 mL, 10 mg mL−1) was added drop by drop to aqueous CCM (5 mL, 2 mg mL−1) under ultrasound. The mixture was then stirred for 24 h and collected by centrifugation at 12,000 rpm for 15 min. The resulting precipitate, referred to as DCCM, was washed three times with water. For the preparation of DCCMH NPs, aqueous HA (1 mL, 10 mg mL−1) was added drop by drop to aqueous DCCM (5 mL, 2 mg mL−1) under ultrasound, the mixture was stirred for 12 h and the precipitate was collected by centrifugation at 12,000 rpm for 15 min. The resulting precipitate, referred to as DCCMH, was washed three times with water.

DOX loading and release from DCCMH NPs

To determine the drug loading efficiency (DLE%) of DCCMH, the centrifugal supernatants obtained during the loading of DOX were collected. The concentration of DOX in the supernatants was measured using a standard curve of DOX determined by UV–Vis spectroscopy. The DLE% and encapsulation efficiency (EE%) of DCCMH were calculated using the following Eqs. (1) and (2). Drug release was assessed under both the physiological and the tumor microenvironments. Specifically, 2 mg of DCCMH NPs were suspended in 4 mL of different phosphate-buffered solutions (pH5.0 with 0 mM GSH, and pH5.0 with 10 mM GSH) and stirred at 37 ℃. At each time interval, 1.5 mL of release medium was extracted to determine the percentage of DOX released through UV–Vis spectrophotometry analysis. The sample was then returned to its original release system for further evaluation.

(1)
(2)

GSH depletion and ·OH generation

The reduction in GSH levels caused by varying concentrations of DCCMH nanoparticles was assessed using the reduced GSH assay kit. Briefly, the 0.5 mL of different concentrations of DCCMH solution were mixed with 0.5 mL of GSH (20 mM) solution, and the mixture was incubated at 37 ℃ for 6 h. Subsequently, the solutions were centrifuged at 12,000 rpm for 10 min, and the absorbance of the supernatant was measured at 412 nm using UV–Vis spectroscopy.

The production of ·OH was measured using MB as a probe. A solution of MB (10 μg mL−1) was added to PBS, H2O2, H2O2 + DCCMH, H2O2 + GSH, and H2O2 + DCCMH + GSH solutions (H2O2: 10 mM, DCCMH: 100 μg mL−1, GSH: 10 mM) and stirred for various durations. The bleaching of MB was monitored by measuring its absorbance at 665 nm using UV–Vis spectroscopy. Additionally, electron spin resonance spectroscopy was employed to verify the generation of ·OH. A mixture of 10 μL of 5,5-dimethyl-1-pyrroline N-oxide (DMPO) and 1 mL of the DCCMH suspension (H2O2: 10 mM, DCCMH: 100 μg mL−1) was measured using an EMX plus spectrometer (Bruker, Germany).

Cell culture and cellular uptake

HL7702 cells, HepG2 cells, Hela cells, U87MG cells, and 4T1 cells were purchased from KeyGEN BioTECH Co. (Nanjing, China). Cells were cultured in high glucose medium DMEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin at 37 ℃ under 5% CO2. For cellular uptake tests, the HepG2 cells were seeded into 6-well plates or CLSM culture dishes and incubated with DCCMH (50 μg mL−1) for different durations (1, 3, 6, and 9 h). Subsequently, the cells were harvested and resuspended in PBS. Flow cytometry was employed to measure the fluorescence intensity of the cells. Additionally, the cells were washed twice with PBS and stained with Hoechst 33258 for 10 min. CLSM was used to capture images of the stained cells (Hoechst 33,258: λex = 405 nm, λem = 460 nm; DOX: λex = 488 nm, λem = 590 nm).

Cytotoxicity assay

Cells were seeded into 96-well plates at a density of 7.0 × 103 cells per well and incubated with various drugs (DOX, CCMH, or DCCMH) for 24 or 48 h. After that, 20 μL of thiazolyl blue tetrazolium bromide (MTT) solution (5 mg mL−1) was added to each well, and cells were further incubated for 4 h. Next, the resulting formazan crystals were dissolved using 100 μL of dimethyl sulfoxide, and the absorbance of each well was measured at 490 nm using a microplate reader (FLUOstar Omega).

Intracellular GSH depletion and ROS generation

HepG2 cells were seeded into T25 culture flasks and allowed to grow for 24 h, then the cells were treated with PBS, DOX (4 μg mL−1), CCMH (46 μg mL−1), and DCCMH (50 μg mL−1), respectively, for 12 h. After removing the culture medium, the cells were collected and subjected to three cycles of freezing and thawing using liquid nitrogen and 37 ℃ water, respectively. The resulting cell lysates were then centrifuged, and the absorbance at 412 nm was measured using a microplate reader (FLUOstar Omega). HepG2 cells were seeded into 6-well plates at a density of 2.0 × 105 cells per well for 24 h.

To measure the level of ROS in the cells, similarly, the cells were cultured with different drugs for 12 h, and after removing the culture medium, 1 mL of DCFH-DA (10 μM) solution was added to each well and cultured for another 30 min. Then, the cells were washed three times with PBS to remove excess DCFH-DA and fluorescence images were captured using CLSM.

In vitro lactic acid and pH measurements

HepG2 cells were seeded into 6-well plates at a density of 2.0 × 105 cells per well and incubated with various drugs (PBS, DOX: 4.0 μg mL−1, CCMH: 46 μg mL−1, DCCMH: 50 μg mL−1) for 12 h, respectively. Then, both the cells and the culture medium were collected for the measurement of lactic acid levels using lactic acid detection kits, following the manufacturer's instructions.

HepG2 cells were cultured in CLSM culture dishes overnight, then the cells were treated with different drugs including PBS, DOX (4.0 μg mL−1), CCMH (46 μg mL−1), and DCCMH (50 μg mL−1) for 12 h. Subsequently, the cells were washed with PBS and incubated with the pH fluorescent probe (BCECF-AM, 5 μM in PBS) for 30 min, and observed using a CLSM. The culture media with a pH of 7.4 was used as standard samples for comparison.

Mitochondrial membrane potential measurement

HepG2 cells were seeded into CLSM culture dishes and cultured overnight. After treatment with PBS, DOX (4.0 μg mL−1), CCMH (46 μg mL−1), and DCCMH (50 μg mL−1) for 12 h, respectively, the cells were incubated with JC-1 dye for 30 min to measure the MMP through CLSM analysis.

Live/dead cell staining and apoptosis assay

HepG2 cells were seeded into CLSM culture dishes with a density of 1.0 × 105 cells per dish for 24 h, and then treated with PBS, DOX (4.0 μg mL−1), CCMH (46 μg mL−1), and DCCMH (50 μg mL−1) for 12 h, respectively. Then, the cells were stained with Calcein-AM (2 μM) and PI (4 μM) for 30 min to measure the cell viabilities through CLSM analysis.

HepG2 cells were cultured with different drugs (PBS, DOX: 4.0 μg mL−1, CCMH: 46 μg mL−1, DCCMH: 50 μg mL−1) for 24 h. Afterward, all of the treated cells were collected, washed, and stained with FITC/PI for 20 min. Then, the intracellular fluorescence signals of FITC/PI were measured by flow cytometry, to determine the apoptosis and necrosis rate of HepG2 cells.

Hemolysis assay

1 mL of red blood cell suspension (0.2%, v/v) was mixed with 1 mL of different concentrations of DCCMH and incubated at 37 ℃ for 4 h. The cells incubated in double distilled water and PBS were used as positive and negative controls, respectively. After centrifugation at 3000 rpm for 6 min, the optical density at 540 nm of each solution was measured using the microplate reader. The hemolysis rate was calculated according to the following formula (3).

(3)

Distribution analysis of DCCMH in vivo

Female BALB/c mice (5–6 weeks old, ~ 18 g each) were inoculated with H22 cells (7.5 × 105 cells per mouse). On day 10, treatment was initiated by intravenous injection of DCCMH-Cy5.5, followed by observation of DCCMH-Cy5.5 distribution within the mice at 0, 2, 4, 6, 8, 12, 24, and 28 h with photographic documentation. Subsequently, the major organs were harvested for ex vivo imaging analysis.

In vivo antitumor evaluation

Female BALB/c mice were subcutaneously inoculated with 0.1 mL of H22 cell suspension (1.0 × 106 cells) into their right leg to establish the tumor model. Subsequently, once the tumor volume reached 100 mm3, the mice were randomly divided into four groups with seven mice in each group: Control (PBS), DOX, CCMH, and DCCMH. The respective formulations were intravenously injected every 4 days for a total of 18 days, with a dose of 0.16 mg kg−1 of DOX, 1.84 mg kg−1 of CCMH, and 2 mg kg−1 of DCCMH. The tumor size and body weight of the mice were measured once every three days. After 18 days of treatment, tumors in all treatment groups were harvested for H&E staining, TUNEL staining, and Ki67 staining. All animal experiments were carried out under the protocols approved by the Institutional Animal Care and Use Committee of Zhejiang University.

Long-term in vivo biosafety evaluation

After the in vivo tumor treatment with different formulations including PBS, DOX (0.16 mg kg−1), CCMH (1.84 mg kg−1), and DCCMH (2 mg kg−1), the major organs (heart, liver, spleen, lungs, and kidneys) of mice were harvested and conducted H&E staining for the histological analysis. The blood samples were collected from the mouse orbital venous plexus for blood biochemistry and blood routine examinations.

Statistical analysis

The one-way analysis of variance (ANOVA) statistical method was performed to evaluate the experimental data. A p-value of 0.05 was selected as the significance level and the data were indicated with (*) for p < 0.05, (**) for p < 0.01, and (***) for p < 0.001, respectively.

Data availability

No datasets were generated or analysed during the current study.

References

  1. Anderson NM, Simon MC. The tumor microenvironment. Curr Biol. 2020;30:921–5.

    Article  Google Scholar 

  2. Siegel RL, Miller KD, Fuchs HE, Jemal A. Cancer statistics, 2022. CA Cancer J Clin. 2022;72:7–33.

    Article  PubMed  Google Scholar 

  3. Di XJ, Pei ZC, Pei YX, James TD. Tumor microenvironment-oriented MOFs for chemodynamic therapy. Coordin Chem Rev. 2023;484: 215098.

    Article  CAS  Google Scholar 

  4. Zhang Y, Zhu JY, Zhang Z, He DN, Zhu J, Chen YS, Zhang YX. Remodeling of tumor microenvironment for enhanced tumor chemodynamic/ photothermal/chemo-therapy. J Nanobiotechnol. 2022;20:388–407.

    Article  Google Scholar 

  5. Sun WJ, Luo L, Feng YS, Qiu YW, Shi CR, Meng SS, Chen XY, Chen HM. Gadolinium–rose bengal coordination polymer nanodots for MR-/fluorescence-image-guided radiation and photodynamic therapy. Adv Mater. 2020;32:2000377.

    Article  CAS  Google Scholar 

  6. Hu FQ, Song D, Yan YM, Huang CS, Shen CT, Lan JQ, Chen YQ, Liu AY, Wu Q, Sun L, Xu F, Hu FY, Chen LS, Luo XL, Feng YD, Huang SY, Hu JB, Wang GH. IL-6 regulates autophagy and chemotherapy resistance by promoting BECN1 phosphorylation. Nat Commun. 2021;12:3651.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Chang YC, Lv YH, Wei P, Zhang PF, Pu L, Chen XX, Yang K, Li XL, Lu YC, Hou CX, Pei YX, Zeng WX, Pei ZC. Multifunctional glyco-nanofibers: siRNA induced supermolecular assembly for codelivery in vivo. Adv Funct Mater. 2017;27:1703083.

    Article  Google Scholar 

  8. Liu X, Ye NB, Liu S, Guan JK, Deng QY, Zhang ZJ, Xiao C, Ding ZY, Zhang BX, Chen XP, Li ZF, Yang XL. Hyperbaric oxygen boosts PD-1 antibody delivery and T cell infiltration for augmented immune responses against solid tumors. Adv Sci. 2021;8:2100233.

    Article  CAS  Google Scholar 

  9. Wang HR, Han X, Dong ZL, Xu J, Wang J, Liu Z. Hyaluronidase with pH-responsive dextran modification as an adjuvant nanomedicine for enhanced photodynamic-immunotherapy of cancer. Adv Funct Mater. 2019;29:1902440.

    Article  Google Scholar 

  10. Kalluri R. The biology and function of fibroblasts in cancer. Nat Rev Cancer. 2016;16:582–98.

    Article  CAS  PubMed  Google Scholar 

  11. Watabe T, Takahashi K, Pietras K, Yoshimatsu Y. Roles of TGF-β signals in tumor microenvironment via regulation of the formation and plasticity of vascular system. Semin Cancer Biol. 2023;92:130–8.

    Article  CAS  PubMed  Google Scholar 

  12. Xiang YT, Chen QH, Nan YY, Lin M, Xiao ZX. Yang YQ, Zhang JP, Ying XH, Long XY, Wang SY, Sun J, Huang Q, Ai KL. Nitric oxide-based nanomedicines for conquering TME fortress: Say “NO” to insufficient tumor treatment. Adv Funct Mater. 2024; 34: 2312092.

  13. Hang YH, Liu YF, Teng ZG, Cao XF, Zhu HT. Mesoporous nanodrug delivery system: a powerful tool for a new paradigm of remodeling of the tumor microenvironment. J Nanobiotechnol. 2023;21:101–25.

    Article  CAS  Google Scholar 

  14. Li JY, Chen Q, Li SY, Zeng XL, Qin JQ, Li X, Chen ZX, Zhang WX, Zhao YB, Huang ZM, Yang XL, Gan L. An adhesive hydrogel implant combining chemotherapy and tumor microenvironment remodeling for preventing postoperative recurrence and metastasis of breast cancer. Chem Eng J. 2023;473: 145212.

    Article  CAS  Google Scholar 

  15. Jalali F, Fakhari F, Sepehr A, Zafari J, Sarajar BO, Sarihi P, Jafarzadeh E. Synergistic anticancer effects of doxorubicin and metformin combination therapy: a systematic review. Transl Oncol. 2024;45: 101946.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Wan MM, Liu ZY, Li T, Chen H, Wang Q, Chen TT, Tao YF, Mao C. Zwitterion-based hydrogen sulfide nanomotors induce multiple acidosis in tumor cells by destroying tumor metabolic symbiosis. Angew Chem Int Ed. 2021;60:16139.

    Article  CAS  Google Scholar 

  17. Yang MY, Li JP, Gu P, Fan XQ. The application of nanoparticles in cancer immunotherapy: targeting tumor microenvironment. Bioact Mater. 2021;6:1973–87.

    CAS  PubMed  Google Scholar 

  18. Calori IR, Piva HL, Tedesco AC. Targeted cancer therapy using alpha-cyano-4-hydroxycinnamic acid as a novel vector molecule: a proof-of-concept study. J Drug Deliv Sci Tec. 2020;57: 101633.

    Article  CAS  Google Scholar 

  19. Zhu HF, Huang DQ, Nie M, Zhao YJ, Sun LY. Dexamethasone loaded DNA scavenger nanogel for systemic lupus erythematosus treatment. Bioact Mater. 2025;43:330–9.

    CAS  Google Scholar 

  20. Dong JL, Yu YY, Pei YX, Pei ZC. pH-responsive aminotriazole doped metal organic frameworks nanoplatform enables self-boosting reactive oxygen species generation through regulating the activity of catalase for targeted chemo/chemodynamic combination therapy. J Colloid Interf Sci. 2022;607:1651–60.

    Article  CAS  Google Scholar 

  21. Shu M, Wang JG, Xu ZY, Lu TL, He Y, Li RS, Zhang GQ, Yan YB, Zhang Y, Chu X, Ke J. Targeting nanoplatform synergistic glutathione depletion-enhanced chemodynamic, microwave dynamic, and selective-microwave thermal to treat lung cancer bone metastasis. Bioact Mater. 2024;39:544–61.

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Sun PF, Qu F, Zhang C, Cheng PF, Li XY, Shen QM, Li DF, Fan QL. NIR-II excitation phototheranostic platform for synergistic photothermal therapy/chemotherapy/chemodynamic therapy of breast cancer bone metastases. Adv Sci. 2022;9:2204718.

    Article  CAS  Google Scholar 

  23. Zhao PR, Li HY, Bu WB. A forward vision for chemodynamic therapy: issues and opportunities. Angew Chem Int Ed. 2023;62: e202210415.

    Article  CAS  Google Scholar 

  24. Peng XX, Zhang H, Zhang RJ, Li ZH, Yang ZS, Zhang J, Gao S, Zhang JL. Gallium triggers ferroptosis through a synergistic mechanism. Angew Chem Int Ed. 2023;62: e202307838.

    Article  CAS  Google Scholar 

  25. Lv MC, Sun M, Wu MC, Zhang F, Yin HY, Sun Y, Liu R, Fan Z, Du JZ. Tryptophan-modulated nanoscale metal–organic framework for coordinated loading of biomolecules for cascade production of reactive oxygen and nitrogen species. Nano Lett. 2022;22:9621–9.

    Article  CAS  PubMed  Google Scholar 

  26. Chen YP, Lyu RY, Wang J, Cheng QC, Yu YF, Yang SX, Mao CB, Yang MY. Metal−organic frameworks nucleated by silk fibroin and modified with tumor-targeting peptides for targeted multimodal cancer therapy. Adv Sci. 2023;10:2302700.

    Article  CAS  Google Scholar 

  27. Feng ZZ, Chen G, Zhong M, Lin L, Mai ZY, Tang Y, Chen GM, Ma W, Li G, Yang YY, Yu ZQ, Yu M. An acid-responsive MOF nanomedicine for augmented anti-tumor immunotherapy via a metal ion interference-mediated pyroptotic pathway. Biomaterials. 2023;302: 122333.

    Article  CAS  PubMed  Google Scholar 

  28. Lu KD, He CB, Guo NN, Chan C, Ni KY, Weichselbaum R, Lin WB. Chlorin-based nanoscale metal–organic framework systemically rejects colorectal cancers via synergistic photodynamic therapy and checkpoint blockade immunotherapy. J Am Chem Soc. 2016;138:12502–10.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Lu KD, Aung T, Guo NN, Weichselbaum R, Lin WB. Nanoscale metal–organic frameworks for therapeutic, imaging, and sensing applications. Adv Mater. 2018;30:1707634.

    Article  Google Scholar 

  30. Luo Y, He XJ, Du QY, Xu L, Xu J, Wang JR, Zhang WL, Zhong YX, Guo DJ, Liu Y, Chen XY. Metal-based smart nanosystems in cancer immunotherapy. Exploration. 2024. https://doiorg.publicaciones.saludcastillayleon.es/10.1002/EXP.20230134.

    Article  PubMed  PubMed Central  Google Scholar 

  31. Du JH, Zhou MT, Chen Q, Tao YC, Ren J, Zhang Y, Qin HL. Disrupting intracellular iron homeostasis by engineered metal-organic framework for nanocatalytic tumor therapy in synergy with autophagy amplification-promoted ferroptosis. Adv Funct Mater. 2023;33:2215244.

    Article  CAS  Google Scholar 

  32. Li YW, Duan YX, Li YY, Gu Y, Zhou L, Xiao ZT, Yu XY, Cai YJ, Cheng EZ, Liu QQ, Jiang Y, Yang Q, Zhang F, Lei Q, Yang B. Cascade loop of ferroptosis induction and immunotherapy based on metal-phenolic networks for combined therapy of colorectal cancer. Exploration. 2024. https://doiorg.publicaciones.saludcastillayleon.es/10.1002/EXP.20230117.

    Article  PubMed  PubMed Central  Google Scholar 

  33. Lu KD, He CB, Guo NN, Chan C, Ni K, Lan GX, Tang HD, Pelizzari C, Fu YX, Spiotto MT, Weichselbaum R, Lin WB. Low-dose X-ray radiotherapy–radiodynamic therapy via nanoscale metal–organic frameworks enhances checkpoint blockade immunotherapy. Nat Biomed Eng. 2018;2:600–10.

    Article  CAS  PubMed  Google Scholar 

  34. Wang WJ, Zhang L, Liu ZQ, Zhang YJ, Zhu JW, Liu MM, Ren JS, Xu XG. Selective methionine pool exhaustion mediated by a sequential positioned MOF nanotransformer for intense cancer immunotherapy. Adv Mater. 2023;35:2211866.

    Article  CAS  Google Scholar 

  35. Han HH, Wang HM, Jangili P, Li ML, Wu LL, Zang Y, Sedgwick AC, Li J, He XP, James TD, Kim JS. The design of small-molecule prodrugs and activatable phototherapeutics for cancer therapy. Chem Soc Rev. 2023;52:879–920.

    Article  CAS  PubMed  Google Scholar 

  36. Lian XZ, Huang YY, Zhu YY, Fang Y, Zhao R, Joseph E, Li JL, Pellois JP, Zhou HC. Enzyme-MOF nanoreactor activates nontoxic paracetamol for cancer therapy. Angew Chem Int Ed. 2018;57:5725–30.

    Article  CAS  Google Scholar 

  37. Yu JY, Li Y, Yan A, Gao YW, Xiao F, Xu ZW, Xu JY, Yu SJ, Liu JQ, Sun HC. Self-propelled enzymatic nanomotors from prodrug-skeletal zeolitic imidazolate frameworks for boosting multimodel cancer therapy efficiency. Adv Sci. 2023;10:2301919.

    Article  CAS  Google Scholar 

  38. Schneider JD, Smith BA, Williams GA, Powell DR, Perez F, Rowe GT, Yang L. Synthesis and characterization of Cu(II) and mixed-valence Cu(I)Cu(II) clusters supported by pyridylamide ligands. Inorg Chem. 2020;59:5433–46.

    Article  CAS  PubMed  Google Scholar 

  39. Sun WJ, Luo L, Feng YS, Cai YT, Zhuang YX, Xie RJ, Chen XY, Chen HM. Aggregation-induced emission gold clustoluminogens for enhanced low-dose X-ray-induced photodynamic therapy. Angew Chem Int Ed. 2020;59:9914–21.

    Article  CAS  Google Scholar 

  40. Attri P, Kim YH, Park DH, Park JH, Hong YJ, Uhm HS, Kim KN, Fridman A, Choi EH. Generation mechanism of hydroxyl radical species and its lifetime prediction during the plasma-initiated ultraviolet (UV) photolysis. Sci Rep. 2015;5:9332–9.

    Article  PubMed  PubMed Central  Google Scholar 

  41. Luo L, Sun WJ, Feng YS, Qin RX, Zhang JH, Ding DD, Shi T, Liu XM, Chen XY, Chen HM. Conjugation of a scintillator complex and gold nanorods for dual-modal image-guided photothermal and X-ray-induced photodynamic therapy of tumors. ACS Appl Mater Interfaces. 2020;12:12591–9.

    Article  CAS  PubMed  Google Scholar 

  42. Dong JL, Ma K, Pei YX, Pei ZC. Core–shell metal–organic frameworks with pH/GSH dual-responsiveness for combined chemo–chemodynamic therapy. Chem Commun. 2022;58:12341–4.

    Article  CAS  Google Scholar 

  43. Zhang X, Wang X, Li Z, Du J, Xiao X, Pan D, Zhang H, Tian X, Gong Q, Gu Z, Luo K. Lactose-modified enzyme-sensitive branched polymers as a nanoscale liver cancer-targeting MRI contrast agent. Nanoscale. 2023;15:809–19.

    Article  PubMed  Google Scholar 

  44. Zhao R, Li T, Zheng G, Jiang K, Fan L, Shao J. Simultaneous inhibition of growth and metastasis of hepatocellular carcinoma by co-delivery of ursolic acid and sorafenib using lactobionic acid modified and pH-sensitive chitosan-conjugated mesoporous silica nanocomplex. Biomaterials. 2017;143:1–16.

    Article  CAS  PubMed  Google Scholar 

  45. Zhao R, Zheng G, Fan L, Shen Z, Jiang K, Guo Y, Shao JW. Carrier-free nanodrug by co-assembly of chemotherapeutic agent and photosensitizer for cancer imaging and chemo-photo combination therapy. Acta Biomater. 2018;70:197–210.

    Article  CAS  PubMed  Google Scholar 

  46. Wan YC, Chen ZL, Wang Y, Zhao WK, Pei ZC, Pu L, Lv YH, Li JX, Li JH, Pei YX. A hyaluronic acid modified cuprous metal-organic complex for reversing multidrug resistance via redox dyshomeostasis. Carbohydr Polym. 2023;311: 120762.

    Article  CAS  PubMed  Google Scholar 

  47. Song MM, Xia WT, Tao ZX, Zhu B, Zhang WX, Liu C, Chen SY. Self-assembled polymeric nanocarrier-mediated co-delivery of metformin and doxorubicin for melanoma therapy. Drug Deliv. 2021;28:594–606.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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Acknowledgements

The authors thank the Teaching and Research Core Facility at the College of Life Science, the Life Science Research Core Services, and the Northwest A&F University for helping with characterizations including SEM, TEM, and CLSM, etc.

Funding

This work was supported by the National Natural Science Foundation of China (22171230, 82001956), the National University of Singapore (NUHSRO/2020/133/Startup/08, NUHSRO/2023/008/NUSMed/TCE/LOA, NUHSRO/2021/034/TRP/09/Nanomedicine, NUHSRO/2021/044/Kickstart/09/LOA, 23-0173-A0001), the National Medical Research Council (MOH-001388-00, CG21APR1005, MOH-001500-00, MOH-001609-00), the Singapore Ministry of Education (MOE-000387-00, MOET32023-0005), the National Research Foundation (NRF-000352-00), the Project of Science and Technology of Social Development in Shaanxi Province (2023YBSF-151 and 2021SF-120), and the 2023 Hangzhou West Lake Pearl Project Leading Innovation Youth Team Project (TD2023017).

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Author notes

  1. Junliang Dong and Jindong Ding have contributed equally to this work.

Authors and Affiliations

  1. College of Chemistry & Pharmacy, Northwest A&F University, Yangling, Shaanxi, 712100, People’s Republic of China

    Junliang Dong, Jindong Ding, Yi Wang, Yuxin Pei & Zhichao Pei

  2. ZJU-Hangzhou Global Scientific and Technological Innovation Center, Xiaoshan District, No.733 Jianshe San Road, Hangzhou, 311200, Zhejiang, People’s Republic of China

    Junliang Dong, Shifan Luo, Ruoshui Li, Yi Wang, Bing Xiao & Wenjing Sun

  3. Department of Diagnostic Radiology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, 119074, Singapore

    Xiaoyuan Chen

  4. Department of Chemical and Biomolecular Engineering, College of Design and Engineering, National University of Singapore, Singapore, 117575, Singapore

    Xiaoyuan Chen

  5. Department of Biomedical Engineering, College of Design and Engineering, National University of Singapore, Singapore, 117575, Singapore

    Xiaoyuan Chen

  6. Department of Pharmacy and Pharmaceutical Sciences, Faculty of Science, National University of Singapore, Singapore, 117544, Singapore

    Xiaoyuan Chen

  7. Clinical Imaging Research Centre, Centre for Translational Medicine, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, 117599, Singapore

    Xiaoyuan Chen

  8. Nanomedicine Translational Research Program, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, 117597, Singapore

    Xiaoyuan Chen

  9. Theranostics Center of Excellence (TCE), Yong Loo Lin School of Medicine, National University of Singapore, Singapore, 138667, Singapore

    Xiaoyuan Chen

  10. Institute of Molecular and Cell Biology, Agency for Science, Technology, and Research (A*STAR), Singapore, 138673, Singapore

    Xiaoyuan Chen

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  1. Junliang Dong
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Contributions

J.D. formulated the conceptual framework and established the experimental procedures, conducted in vitro and in vivo experiments, analyzed data, drafted the manuscript, compiled all figures, and revised the manuscript based on the feedback from W.S., X.C., and Z.P.. J.D. assisted in writing and revising the manuscript. S.L., R.L., Y.W., and B.X. participated in conducting the research. Y.P., W.S., X.C., and Z.P. collaboratively reviewed and edited the manuscript, and provided financial support. All authors collectively analyzed the findings, revised the manuscript, and unanimously endorsed the final version.

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Correspondence to Yuxin Pei, Xiaoyuan Chen, Wenjing Sun or Zhichao Pei.

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Dong, J., Ding, J., Luo, S. et al. Remodeling tumor microenvironment using prodrug nMOFs for synergistic cancer therapy. J Nanobiotechnol 23, 123 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12951-025-03202-7

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