Abstract Background Radiotherapy (RT) is a primary clinical approach for cancer treatment, but its efficacy is often hindered by various challenges, especially radiation resistance, which greatly compromises the therapeutic effectiveness of RT. Mitochondria, central to cellular energy metabolism and regulation of cell death, play a critical role in mechanisms of radioresistance. In this context, cuproptosis, a novel copper-induced mitochondria-respiratory-dependent cell death pathway, offers a promising avenue for radiosensitization. Results In this study, an innovative theranostic nanoplatform was designed to induce cuproptosis in synergy with low-dose radiation therapy (LDRT, i.e., 0.5–2 Gy) for the treatment of in situ hepatocellular carcinoma (HCC). This approach aims to reverse the hypoxic tumor microenvironment, promoting a shift in cellular metabolism from glycolysis to oxidative phosphorylation (OXPHOS), thereby enhancing sensitivity to cuproptosis. Concurrently, the Fenton-like reaction ensures a sustained supply of copper and depletion of glutathione (GSH), inducing cuproptosis, disrupting mitochondrial function, and interrupting the energy supply. This strategy effectively overcomes radioresistance and enhances the therapeutic efficacy against tumors. Conclusions In conclusion, this study elucidates the intricate interactions among tumor hypoxia reversal, cuproptosis, metabolic reprogramming, and radiosensitization, particularly in the context of treating in situ hepatocellular carcinoma, thereby providing a novel paradigm for radiotherapy. Graphical abstract [42]graphic file with name 12951_2024_3011_Figa_HTML.jpg Supplementary Information The online version contains supplementary material available at 10.1186/s12951-024-03011-4. Keywords: Cuproptosis, Low-dose radiation therapy, Metabolic reprogramming, Radiosensitization, Mitochondria Background Cancer continues to be a leading cause of global mortality, propelling continuous exploration into various therapeutic modalities to mitigate its impact [[43]1, [44]2]. Radiotherapy, as an effective non-invasive approach, inflicts damage on biomacromolecules such as DNA, proteins, and lipids through direct or indirect effects, leading to tumor cell death [[45]3, [46]4]. Despite over 50% of cancer patients undergoing radiotherapy, only a small fraction exhibit complete treatment responses [[47]5, [48]6]. This is primarily due to the radioresistance of tumor tissues and the maximum tolerable dose limitations of adjacent normal tissues [[49]7, [50]8]. Radioresistance is a complex process involving alterations in multiple cellular mechanisms [[51]9]. Notably, altered mitochondria morphology and metabolism induced by tumors contribute to radioresistance through energy metabolism reprogramming and apoptosis evasion, making mitochondria a potential target for radiosensitization [[52]10, [53]11]. Mitochondrial metabolic reprogramming, one of the major factors of radioresistance, enables tumor cells to rapidly adapt to hypoxic and nutrient-deprived tumor microenvironment by shifting from OXPHOS to glycolysis, thus promoting rapid proliferation [[54]12–[55]17]. Consequently, reprogramming metabolism and cutting off the energy supply is crucial for effective cancer treatment [[56]18–[57]21]. On the other hand, mitochondria, regulate cell death through various mechanisms such as apoptosis, autophagy, and necrosis [[58]22, [59]23]. However, tumor-induced disorder of apoptosis signaling, undermines the efficacy of radiotherapy and contributes to radioresistance [[60]24]. Recently, Tsvetkov et al. discovered a new mitochondria-respiratory-dependent cell death mechanism induced by copper, termed cuproptosis, opening up new avenues for utilizing copper in cancer therapy [[61]25]. Copper, as a crucial intracellular trace element, is involved in nearly all fundamental life processes [[62]26]. Recent studies have found that excess copper within cells directly binds to dihydrolipoamide S-acetyltransferase (DLAT) of the tricarboxylic acid cycle (TCA), inducing the oligomerization of acetylated DLAT and disrupting OXPHOS metabolism [[63]27]. Simultaneously, ferredoxin (FDX1) facilitates the reduction of Cu(II) to the more toxic Cu(I), further leading to protein toxicity stress and inducing cell death [[64]28, [65]29]. Cuproptosis-induced mitochondrial damage also reduces intermediates in the TCA, impairing cell proliferation and DNA damage repair capabilities, which is also expected to increase radiosensitivity. Notably, cuproptosis is a mitochondria-respiratory-dependent cell death mode, which implies that the hypoxic tumor microenvironment within solid tumors significantly affects their sensitivity to cuproptosis [[66]27, [67]30, [68]31]. Additionally, the high concentration of GSH within tumor cells, acting as a thiol-containing copper chelator, can inhibit the process of cuproptosis [[69]32–[70]34]. Addressing these challenges is crucial for achieving radiosensitization through cuproptosis in clinic. We have successfully developed a theranostic nanoplatform (HA-PEG@CuO[2]) that harnesses Fenton-like reactions and cuproptosis, providing a multifaceted approach to tumor treatment by alleviating hypoxia, reprogramming metabolism, and inducing mitochondrial damage. Radiotherapy remains a cornerstone treatment for metastatic or unresectable HCC, a typically hypoxic tumor [[71]8, [72]35]. Thus, we established an in situ HCC mouse model and evaluated the antitumor effects of HA-PEG@CuO[2] in combination with LDRT, which significantly minimizes damage to normal tissues [[73]36]. Notably, HA-PEG@CuO[2] demonstrates several advantages: (1) Mediating Fenton-like reactions: HA-PEG@CuO[2], with excellent targeting and tumor microenvironment responsiveness, undergoes Fenton-like reactions to generate highly active •OH radicals and deplete GSH, increasing sensitivity to cuproptosis. (2) Alleviating Tumor Hypoxia: CuO[2] produces O[2], effectively reversing the hypoxic tumor microenvironment, which in turn regulates tumor metabolism and increases sensitivity to cuproptosis. (3) Inducing Cuproptosis: Sustained release of copper ions facilitates the direct binding of copper to lipidated TCA cycle components, triggering cuproptosis. (4) Bimodal Imaging: The inherent T1 enhancement properties of Cu(II) and the near-infrared fluorescence imaging capabilities of ICG endow HA-PEG@CuO[2] with excellent dual-modal imaging capabilities. In summary, this study provides a novel strategy for LDRT by reversing the hypoxic tumor microenvironment and regulating tumor metabolism to enhance cuproptosis and mitochondrial damage. Materials and methods Materials Polyvinylpyrrolidone (PVP), hyaluronic acid (HA), 5,5′-Dithiobis-(2-nitrobenzoic acid) (DTNB), and methylene blue (MB) were purchased from Meryer (Shanghai, China). CuCl[2] and NaOH were purchased from Macklin (Shanghai, China). Primary antibodies (11577-1-AP LIAS Rabbit Polyclonal Antibody, 12592-1-AP FDX1 Rabbit Polyclonal Antibody, 13426-1-AP DLAT Rabbit Polyclonal Antibody, CD44 Antibody, and anti-HIF-1α) were obtained from either Affinity (USA) or Proteintech (USA). Ceturegel^® Matrix High Concentration, LDEV-Free was purchased from YEASEN (Shanghai, China). D-Luciferin-potassium was purchased from Meilunbio (Daliang, China). Reactive Oxygen Species Assay Kit, DNA Damage Assay Kit by γ-H[2]AX Immunofluorescence, Mitochondrial membrane potential assay kit with JC-1, Hoechst 33342, Cell Counting Kit-8, and DAPI Staining Solution were purchased from Beyotime (Shanghai, China). Calcein-AM/PI Double Staining Kit was obtained from KEYGEN BIOTECH (Jiangsu, China). Lactic Acid (LA) Content Assay Kit was obtained from Solarbio (Beijing, China). Preparation and characterization of HA-PEG@CuO[2] A 5 mL solution of PVP at a concentration of 0.1 g/mL was prepared. This solution was then mixed with 4 mL of deionized water. Under gentle stirring, 0.2 M CuCl[2] (0.4 mL) and 0.2 M NaOH (0.8 mL) were added to the mixture. After 10 min of stirring at room temperature, 150 μL of 30% hydrogen peroxide (H[2]O[2]) was added dropwise, followed by continuous stirring for 30–60 min. The resulting mixture was centrifuged at 12,000 rpm for 15 min to remove the supernatant. The precipitate was collected and then separately mixed with 10 mg of polyethylene glycol disulfide (PEG-SS-PEG) and 3–5 mg of hyaluronic acid (HA). Subsequently, 10 mL of deionized water was added, and the mixture was stirred for an additional 30–60 min, followed by centrifugation to collect the precipitate. The morphology and elemental distribution of HA-PEG@CuO[2] were characterized by high-resolution transmission electron microscopy (HR-TEM, JEOL 2010, 200 kV) and energy dispersive X-ray spectroscopy (EDS, X-MaxTEM). Particle size and zeta potential of HA-PEG@CuO[2] were analyzed by Zetasizer Nano ZS analyzer (Malvern Instruments Limited). The chemical structures of HA-PEG@CuO[2] were analyzed by inductively coupled plasma mass (ICP-MS, Optima 2000 DV), UV–vis–NIR spectrophotometry (Carry 5000 spectrophotometer), FTIR spectroscopy (Nicolet iS10, Thermo Fisher Scientific), X-ray photoelectron spectroscopy (XPS, Thermo Scientific Escalab 250), 3.0 T MRI scanner (GE 3.0 T), Raman (Renishaw inVia), and NMR (Bruker-600 M Hz NMR). Hemolysis assay Mice-derived red blood cells (RBCs) underwent washing with saline followed by centrifugation until the supernatant clarified for a hemolytic assessment. Deionized water was used as the positive control, while PBS served as the negative control. RBCs were incubated with various concentrations of HA-PEG@CuO[2] at room temperature for 4 h, followed by vortexing and centrifugation at 3000 rpm for 10 min. Subsequently, the supernatant underwent collection, and the optical density (OD) at 545 nm was measured using a microplate reader to determine the hemolysis ratio. And the morphology of red blood cells was observed under the microscope. Cell lines and cell culture SMMC 7721, Hep G2 and LO2 cells were obtained from the Cell Bank of the Chinese Academy of Sciences (China). SMMC 7721 and Hep G2 cells were incubated in DMEM medium (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% FBS. LO2 cells were incubated in RPMI-1640 medium (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% FBS (HyClone, GE Healthcare). They were both incubated in a controlled environment with 5% CO[2] at 37 °C. Cytotoxicity assessment in vitro For the cytotoxicity assessment in vitro, SMMC 7721 cells were seeded in 96-well plates and cultured overnight at 37 °C. Subsequently, the original medium was substituted with fresh medium containing varied concentrations of test samples (PBS, CuCl[2], PVP@CuO[2] or HA-PEG@CuO[2]). After incubation for another 12 h, the cells were then irradiated with or without a radiation dose of 2 Gy and then incubated for another 24 h. Cell viability was assessed using the CCK-8 assay with a microplate reader (Multiskan GO, Thermo Scientific, USA) set to measure absorbance at 450 nm. Cell viability (%) was determined by comparing the absorbance of treated cells to that of untreated cells. Additionally, cytotoxicity was further evaluated using live/dead staining and the colony formation assay. Fluorescence imaging For cellular uptake evaluation, SMMC 7721 and Hep G2 cells were initially seeded in confocal dish and incubated with FITC-HA-PEG@CuO[2] for 0, 2 h, 4 h and 8 h. Additionally, SMMC 7721 cells were subjected to pre-treatment with an excess amount of hyaluronic acid before being co-cultured with FITC-HA-PEG@CuO[2] for the same durations. Finally, intracellular fluorescence was observed by Confocal laser scanning microscope (CLSM, Leica/TCS SP8). For mitochondrial membrane potential evaluation, SMMC 7721 cells were seeded in confocal dish and incubated with PBS, CuCl[2] or HA-PEG@CuO[2], either alone or in combination with a radiation dose of 2 Gy. Afterwards, the medium was aspirated and replaced with 1 ml of JC-1 staining solution. Following thorough mixing, the plates were placed in a cell incubator at 37 °C for a duration of 20 min. Subsequently, the supernatants were carefully aspirated, and the cells were washed twice with JC-1 staining buffer. CLSM was then used for observation. For γ-H[2]AX evaluation, SMMC 7721 cells treated with the above-mentioned formulations, were then stained with γ-H[2]AX antibodies, and imaging was performed using CLSM. For ROS levels evaluation, SMMC 7721 cells treated with the above-mentioned formulations, were stained using a kit designed for detecting reactive oxygen species, and their fluorescence was visualized using CLSM. Cellular GSH/GSSG detection SMMC 7721 cells were seeded in 6-well plates and were treated with different concentrations of HA-PEG@CuO[2] for 24 h. Afterward, the medium was discarded and washed with PBS. The cells were then detached by scraping, followed by centrifugation to eliminate the supernatant and collect the cellular precipitate. This precipitate underwent two rapid freeze–thaw cycles using liquid nitrogen and a water bath set at 37 °C. Finally, the supernatant was obtained via centrifugation after a 5-min incubation in an ice bath and utilized for GSH and GSSG determination employing assay kits. Morphological observation of mitochondria The SMMC 7721 cells were seeded in 6-well plate incubated with PBS, CuCl[2] or HA-PEG@CuO[2], either alone or in combination with a radiation dose of 2 Gy. Afterward, the cells were harvested and fixed with 2.5% glutaraldehyde. Subsequently, the cells were sliced using a slicer to facilitate the acquisition of bio-TEM images. Cellular ATP detection SMMC 7721 cells were seeded in 6-well plates and incubated with PBS, CuCl[2] or HA-PEG@CuO[2], either alone or in combination with a radiation dose of 2 Gy. After collecting the cell precipitate, the cells were fully lysed by adding lysis solution. Subsequently, the supernatant was extracted via centrifugation post-lysis and subjected to analysis utilizing an ATP assay kit. Cellular lactate detection The SMMC 7721 cells were seeded in 6-well plates and incubated with PBS, CuCl[2] or HA-PEG@CuO[2], either alone or in combination with a radiation dose of 2 Gy. Lactate production in cells was assessed using Lactic Acid (LA) Content Assay Kit. This assay enabled the quantification of lactate by measuring the absorbance at 570 nm. Western blotting For western blot (WB) analysis, SMMC 7721 cells were seeded in 6-well plates and incubated with PBS, CuCl[2] and HA-PEG@CuO[2], either alone or in combination with a radiation dose of 2 Gy. Following incubation, the cells were lysed on ice using a lysis buffer to extract proteins for sample preparation. Protein content variations, including DLAT oligomers, LIAS, FDX1, and HIF-1α, were then analyzed via gel electrophoresis and antibody detection, culminating in protein visualization. Flow cytometric analysis SMMC 7721 cells pre-seeded in 6-well plates were treated with different methods. After rinsing with PBS, the cell sediment was collected. Flow cytometry analysis was carried out using FLOWJO 10.7.1 software to further examine the cells. Animal tumor models construction and anti-tumor therapy efficacy in vivo Male BALB/c nude mice aged 4–5 weeks were procured from Zhuhai BesTest Bio-Tech Co,.Ltd. (Zhuhai, China). To construct the subcutaneous tumor mice model, 100 μL of SMMMC 7721 cell suspension (1 × 10^6 cells/mL) was subcutaneously injected into each mouse; To construct the in situ HCC mouse model, the mouse liver was exposed, and SMMC 7721 cells were inoculated into the liver by puncturing the left or middle lobe with an 18 G trocar. The abdomen was then closed with absorbable sutures. To ensure the uniformity of tumor formation, all procedures were performed by the same operator. The animal experiments were conducted in accordance with the protocol approved by the Ministry of Health in People’s Republic of PR China (document no. 55, 2001) and the Laboratory Animal Welfare and Ethics Committee of Jinan University (IACUC-20240104-06). Following successful model establishment, mice were randomly divided into six groups: (1) Control (PBS); (2) CuCl[2]; (3) HA-PEG@CuO[2]; (4) X-ray; (5) CuCl[2] + X-ray; (6) HA-PEG@CuO[2] + X-ray. On days 1, 6, and 11, mice were administered 100 μL of the respective nanomaterials (4 mg/kg), with radiotherapy (2 Gy) conducted on days 2, 7 and 12, and subsequent monitoring of mice body weight every day until the end of the experiment. Concurrently, tumor growth was monitored on days 0, 5, 10, 15, and 20 using MRI and FI, providing visual representation of changes in tumor volume during the treatment period. To measure the tumor length and tumor width and the tumor volume was calculated according to the formula. Tumor volume = tumor length × tumor width^2/2. Finally, upon euthanasia, tumors and liver tissues were fixed in 4% neutral buffered formalin, processed routinely into paraffin, and sectioned at a thickness of 4 μm. Then the sections were stained with DLAT, FDX1, LIAS, HIF-1α, ROS and γ-H[2]AX. In vivo magnetic resonance imaging and fluorescence imaging of the biodistribution of HA-PEG@CuO[2] For the biodistribution study, ICG-labeled HA-PEG@CuO[2] (IHA-PEG@CuO[2]) (4 mg/kg) or equal concentration of CuCl[2] was administered into tumor-bearing mice by intravenous injection. In vivo images were recorded using the MRI and IVIS systems at 0, 1, 2, 4, 8, 12 and 24 h post-injection. Mice were sacrificed 24 h after injection to collect the tumors and major organs for imaging analysis using the IVIS system. Biosafety evaluation of HA-PEG@CuO[2] For the biosafety study, male balb/c nude mice were used to evaluate the biosafety of HA-PEG@CuO[2]. Mice were intravenously injected with 100 µL CuCl[2] or HA-PEG@CuO[2] (4 mg/kg). Following injection, blood samples were obtained and subjected to centrifugation at 3000 rpm to isolate serum samples. Biochemical indices in serum, including WBC, HCT, MCV, HGB, RBC, PLT, ALT, AST and CR. Hearts, livers, spleens, lungs, and kidneys from different treatment groups were fixed in 4% paraformaldehyde. Subsequently, they were embedded in paraffin, sectioned, rehydrated, and subjected to hematoxylin and eosin staining. Statistical analysis Data analyses and figures were conducted using the SPSS 24.0 and GraphPad Prism 5.0 software. All values are expressed as mean ± standard deviation (SD). Statistical significance between the groups was determined by t-test or one-way analysis of variance (ANOVA). *P < 0.05, **P < 0.01, ***P < 0.001. Results Preparation and characterization of HA-PEG@CuO[2] Referring to the literature, Polyvinylpyrrolidone-coated copper peroxide nanodots (PVP@CuO[2]) have been successfully synthesized [[74]37] (Fig. [75]1A). Then, the final product HA-PEG@CuO[2] with satisfactory tumor targeting capability and tumor microenvironment responsivity was obtained by simply mixing PVP@CuO[2] with hyaluronic acid (HA) and PEG-SS-PEG. Dynamic light scattering (DLS) data indicate that (Fig. [76]1B and Figure S1, Supporting Information), compared to PVP@CuO[2] (351.7 nm, PDI = 0.551), the modified HA-PEG@CuO[2] exhibits superior dispersibility and a smaller hydrodynamic diameter (162.9 nm, PDI = 0.1067). Additionally, transmission electron microscopy(TEM) images confirm that the morphology of PVP@CuO[2] and HA-PEG@CuO[2] were fusiform-shape with uniform size distribution (Fig. [77]1D). Further confirmation of the presence of clear elemental signals of C, O, N, Cl, and Cu in HA-PEG@CuO[2] was obtained through elemental mapping (Fig. [78]1E). Furthermore, zeta potential measurements revealed that HA-PEG@CuO[2] exhibited a stronger negative zeta potential after modification, which would be beneficial for prolonged circulation in vivo (Fig. [79]1C). The excellent stability and low systemic toxicity of materials are crucial for drug preservation and clinical translation. Thus, HA-PEG@CuO[2] and PVP@CuO[2] were dispersed in phosphate buffered saline (PBS) and DMEM with 10% FBS, and the stability of the nanomaterials was continuously monitored. The results indicated that HA-PEG@CuO[2] remained well-dispersed without obvious size changes during a 72-h storage period (Figure S2, Supporting Information). In addition, the hemocompatibility of HA-PEG@CuO[2] was further validated through hemolysis assays, using water as the positive control and PBS as the negative control. The results demonstrated that even at a drug concentration of 200 μg/ml, the hemolysis rate remained well below the permissible limit (5%) (Figure S3A, Supporting Information). Furthermore, the morphology of red blood cells (RBCs), as observed under an optical microscope, corroborated the hemolysis results. After incubation with HA-PEG@CuO[2], the RBCs retained their original morphology, similar to the negative control (PBS) (Figure S3B, Supporting Information). The presence of CuO[2] within the HA-PEG@CuO[2]was confirmed via ultraviolet–visible (UV–Vis) absorption spectroscopy (Fig. [80]1F). The degree of modification with HA was further quantified to be approximately 12.8%, expressed as the weight percentage of HA. Additionally, the characteristic peaks of all components were evident in both the ^1H-NMR and Fourier-transform infrared (FTIR) spectra of HA-PEG@CuO[2] (Fig. [81]1G and H). In the FTIR spectrum, the peaks at 412 cm^−1 and 520 cm^−1 corresponded to the Cu = O and Cu–O of CuO[2]. The other peak at 1662 cm^−1 was attributed to amide bonding, and the above proved that the successful synthesis of HA-PEG@CuO[2]. Fig. 1. [82]Fig. 1 [83]Open in a new tab Preparation and Characterization of HA-PEG@CuO[2]. A Schematic illustration of the synthesis of the PVP@CuO[2] and HA-PEG@CuO[2]. B Particle size and C Zeta potential of PVP@CuO[2] and HA-PEG@CuO[2]. D TEM image of HA-PEG@CuO[2]. Scale bars = 500.0 nm and 200.0 nm. E Compositional elemental mapping of HA-PEG@CuO[2]. Scale bar = 100 nm. F UV–vis absorption spectrum of CuO[2] and HA-PEG@CuO[2]. G ^1H-NMR spectra of Cystamine, PEG, and HA-PEG@CuO[2]. H FTIR spectra of HA, CuO[2], and HA-PEG@CuO[2]. Data are shown as mean ± SD (n = 3) The fenton-like properties of HA-PEG@CuO[2] caused by pH and GSH dual-responsive Weak acidity, endogenous GSH, and hypoxia are widely recognized characteristics of most tumor microenvironments (TMEs) [[84]38, [85]39]. Thus, the pH and GSH dual-responsive degradation behavior is crucial for mediating the release of tumor microenvironment-responsive drugs and ensuring precise drug delivery. Based on the chemical instability of CuO[2], HA-PEG@CuO[2] underwent the Fenton-like reaction in the tumor microenvironment (as illustrated in Fig. [86]2A), leading to the depletion of GSH and the generation of ·OH. First, to further clarify the chemical structural changes of HA-PEG@CuO[2] under different conditions, Raman spectroscopy and ^1H-NMR were performed. As depicted in Fig. [87]2B, when treated with 10 mM GSH (pH 6.8), the disulfide bonds on the surface of HA-PEG@CuO[2] were cleaved, leading to the release of CuO[2] and a reduction in the characteristic disulfide bond peaks. Furthermore, the ^1H-NMR results corroborated the GSH responsiveness of HA-PEG@CuO[2]. Upon incubation with 10 mM GSH (pH 6.8), the appearance of the chemical shift of -SH (δ = 2.05) confirmed the reduction of disulfide bonds (S–S) to thiol groups (–SH) by GSH (Fig. [88]2C). Subsequently, the full X-ray photoelectron spectroscopy (XPS) spectra revealed the presence of C, O, N and Cu in HA-PEG@CuO[2] both before and after the reaction (Figure S4, Supporting Information). Additionally, high-resolution XPS analysis of Cu 2p confirmed the valence state of Cu in HA-PEG@CuO[2] was + 2, based on the Cu 2p XPS spectra displaying two main peaks at 934.16/953.96 eV, accompanied by two satellite peaks at 942.31/962.05 eV, respectively (Fig. [89]2D). After reaction, the original satellite peak nearly disappeared, and the binding energy peaks shifted from 934.16/953.96 eV to 932.34/952.09 eV, further confirming changes in the oxidation state of copper ions (Figure S5, Supporting Information). Additionally, the O 1 s peaks at 531.16 eV and 532.71 eV were attributed to C = O and O − O, respectively, indicating the presence of HA and peroxy groups. Notably, the intensity of the O–O peak significantly diminishes post-reaction, indicating alterations in the O–O bond (Fig. [90]2E and F). Subsequently, we further evaluated the responsivity of HA-PEG@CuO[2] at different pH (6.8 and 7.4), with or without radiotherapy by assessing the cumulative release of Cu(II) (Fig. [91]2G and Figure S6, Supporting Information). As depicted in Fig. [92]2G, HA-PEG@CuO[2] exhibited exceptional chemical stability at pH 7.4 compared to PH 6.8, with a mere 8.42% release rate of Cu(II) after 48 h. Furthermore, we also revealed that the presence of GSH significantly accelerated the release rate of Cu(II) at both pH values, which could be attributed to GSH-induced disulfide bond cleavage. For example, under conditions of pH = 6.8 (10 mM GSH), HA-PEG@CuO[2] demonstrated a pronounced burst release behavior, with the release rate of Cu(II) increased up to 76.50% after 48 h, markedly higher than the release rate of 13.23% observed at the same pH without GSH. Fig. 2. [93]Fig. 2 [94]Open in a new tab The Fenton-like Properties of HA-PEG@CuO[2] caused by pH and GSH Dual-Responsive. A Schematic illustration of the pH and GSH dual-responsive and Fenton-like properties of HA-PEG@CuO[2]. B Raman spectrum of HA-PEG@CuO[2] and HA-PEG@CuO[2] + GSH. C ^1H-NMR absorption spectrum of HA-PEG@CuO[2] and HA-PEG@CuO[2] + GSH. D Cu2p and E O1s spectrum of HA-PEG@CuO[2]. F O1s spectrum of HA-PEG@CuO[2] + GSH. G Cumulative release profiles of Cu(II) from HA-PEG@CuO[2] dispersed in different pH buffers with and without 10 mM GSH. H UV–vis absorbance (325 nm and 412 nm) of DTNB solution after treated with HA-PEG@CuO[2] in different concentrations (0 μg/mL, 4 μg/mL, 20 μg/mL and 40 μg/mL). I UV–Vis absorbance (665 nm) of MB solution after treated with HA-PEG@CuO[2] dispersed in different pH buffers with and without 10 mM GSH. (1: MB; 2: MB + CuCl[2]; 3: MB + HA-PEG@CuO[2]; 4: pH = 6.8 MB + HA-PEG@CuO[2]; 5: pH = 7.4 MB + HA-PEG@CuO[2]). J UV–vis absorbance (665 nm) of MB solution after treated with HA-PEG@CuO[2] in different concentrations (1: HA-PEG@CuO[2] (0 μg/mL) + GSH; 2: HA-PEG@CuO[2] (4 μg/mL) + GSH; 3: HA-PEG@CuO[2] (20 μg/mL) + GSH; 4: HA-PEG@CuO[2] (40 μg/mL) + GSH). Data are shown as mean ± SD (n = 3) To further validate the Fenton-like catalytic activity of HA-PEG@CuO[2], we conducted further experiments to measure the consumption of GSH and the generation of ·OH. Tumor cells contain abundant GSH, which not only can chelate copper to inhibit cuproptosis but also maintain cellular redox balance and scavenge ROS [[95]40]. Consequently, the rapid depletion of GSH can disrupt redox homeostasis within tumor cells and enhance the sensitivity of cuproptosis and radiotherapy [[96]41, [97]42]. Thus, the GSH-depleting ability of HA-PEG@CuO[2] was evaluated using 5,5′-Dithiobis-(2-nitrobenzoic acid) (DTNB), as a GSH indicator. DTNB exhibits a characteristic absorption peak at 325 nm, which reacts with GSH to produce yellow 2-nitro-5-thiobenzoic acid (TNB) with a typical absorption peak at 412 nm. As shown in Fig. [98]2H, with increasing concentrations of HA-PEG@CuO[2], the absorbance at 412 nm significantly decreased indicating effective GSH consumption. Moreover, the reaction of HA-PEG@CuO[2] with GSH generates Cu(I), which catalyzed the Fenton-like reaction to produce highly oxidative ·OH from endogenous hydrogen peroxide, damaging tumor cell DNA. Using methylene blue (MB) as a probe, the catalytic activity of HA-PEG@CuO[2] in generating ·OH from H[2]O[2] was investigated. MB, sensitive to ·OH, undergoes oxidation, leading to a decrease in absorbance at 665 nm. As shown in Fig. [99]2I, after incubation of HA-PEG@CuO[2] with GSH (pH 6.8), a notable fading of the MB solution was observed, which was corroborated by a decrease in absorbance at 665 nm. Other groups did not observe significant fading of MB. In addition, the absorbance of MB decreased as the concentration of HA-PEG@CuO[2] increased (Fig. [100]2J), confirming abundant generation of ·OH. These results collectively demonstrated that HA-PEG@CuO[2] possessed superior tumor microenvironment responsiveness and Fenton-like catalytic activity. In vitro cellular uptake and cytotoxicity The cluster of differentiation 44 (CD44), a transmembrane glycoprotein receptor, is specifically targeted by HA and is frequently overexpressed on the surface of various cancer cell types [[101]43, [102]44]. This overexpression is closely associated with tumor metastasis, invasion, and prognosis [[103]45, [104]46]. Consequently, HA-modified HA-PEG@CuO[2] can specifically target tumor cells. Initially, WB experiments were conducted to detect the expression of CD44 on the surfaces of LO2 (human normal liver cells), Hep G2 (human liver cancer cells), and SMMC 7721 (human liver cancer cells). The results indicated significantly higher CD44 expression in SMMC 7721 (Fig. [105]3C). Subsequently, to further confirm the targeting specificity and cellular uptake ability of HA-PEG@CuO[2], FITC-labeled HA-PEG@CuO[2] was separately incubated with Hep G2 and SMMC 7721 cells. CLSM results intuitively proved that the fluorescence intensity within SMMC 7721 cells was markedly higher than that within Hep G2 cells (Fig. [106]3A and Figure S7, Supporting Information). Moreover, the fluorescence intensity increased significantly with prolonged incubation time, indicating sustained internalization of HA-PEG@CuO[2] (Fig. [107]3B). Similar results were obtained through quantitative analysis using flow cytometry (Fig. [108]3D). Furthermore, the competitive binding experiment further revealed the targeting specificity of HA-PEG@CuO[2]. Preincubation with an excess of HA for CD44 blockade significantly inhibited cellular uptake of HA-PEG@CuO[2]. As expected, HA-PEG@CuO[2] selectively recognized tumor cells with high CD44 expression and promoted their preferential internalization into tumor cells through CD44 receptor-mediated endocytosis, demonstrating potential for targeted delivery. Fig. 3. [109]Fig. 3 [110]Open in a new tab Effects of HA-PEG@CuO[2] on cellular uptake, cytotoxicity and fenton-like properties in vitro. A CLSM images and B corresponding fluorescence intensities of Hep G2 and SMMC 7721 cells after treatment with FITC-HA-PEG@CuO[2] or FITC-HA-PEG@CuO[2] combined with 1 mg/mL free hyaluronic acid for 8 h. Scale bar = 100 µm. (Blue: DAPI; Green: FITC). C Western blot analysis of CD44 (82 kD) expression in LO2, Hep G2 and SMMC 7721 cells. D Flow cytometric analysis of SMMC 7721 cells incubated with FITC-HA-PEG@CuO[2] at different concentrations. E Cytotoxicity of CuCl[2], PVP@CuO[2] and HA-PEG@CuO[2] either individually or combined with X-ray against SMMC 7721 cells. F Live/dead cell staining images and corresponding fluorescence intensities of SMMC 7721 cells incubated with 40 µM of CuCl[2,] PVP@CuO[2], and HA-PEG@CuO[2] either individually or combined with X-ray. Scale bar = 100 µm (Red: dead cells; Green: living cells). G Representative cloning evaluation images of tumor cells incubated with 40 µM of CuCl[2], PVP@CuO[2] and HA-PEG@CuO[2] either individually or combined with X-ray. H GSH and GSSG levels in cells incubated with HA-PEG@CuO[2] at different concentrations. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA. (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) Subsequently, to validate the in vitro anti-tumor and radiosensitization effects of HA-PEG@CuO[2], different formulations were first assessed for their cytotoxicity against SMMC 7721 cells using the Cell Counting Kit-8 (CCK-8) assay. As shown in Fig. [111]3E, Figure S8 and Table S1, Supporting Information, HA-PEG@CuO[2] exhibited higher cytotoxicity against SMMC 7721 cells (IC50 = 84.42 µM) compared to free CuCl[2] (IC50 = 210.20 µM) and PVP@CuO[2] (IC50 = 108.10 µM). Furthermore, the synergistic effect of HA-PEG@CuO[2] combined with radiotherapy was evaluated. As expected, HA-PEG@CuO[2] demonstrated enhanced cytotoxicity (IC50 = 23.15 µM) when combined with radiotherapy, surpassing the effects of PVP@CuO[2] (IC50 = 67.20 µM). To further verify the radiotherapy sensitization effect of HA-PEG@CuO[2], we conducted live/dead staining and clone formation assay to gain a more intuitive understanding of the cellular viability. Results indicated that the combination of HA-PEG@CuO[2] with radiotherapy exhibited the least amount of cell viability and colony formation (Fig. [112]3F, G and Figure S9, Supporting Information). Overall, HA-PEG@CuO[2] exhibited promising potential for targeted delivery to CD44-overexpressing tumor cells, as well as enhanced efficacy in combination with radiotherapy. To further demonstrate that HA-PEG@CuO[2] could mediate a potent Fenton-like reaction to deplete GSH, an endogenous antioxidative substance and Cu-chelating agent, we investigated the GSH scavenging capability of HA-PEG@CuO[2] at the cellular level. As illustrated in Fig. [113]3H, there was a dose-dependent decrease in the intracellular GSH/GSSG ratio with increasing concentrations of HA-PEG@CuO[2]. This indicated that HA-PEG@CuO[2], once internalized by the cells, underwent a Fenton-like reaction, oxidizing GSH to GSSG. Mechanistic insights into cuproptosis and radiosensitization induced by HA-PEG@CuO[2] Recent studies have indicated that cuproptosis is mediated by the direct binding of copper to DLAT, inducing aberrant oligomerization of lipoylated DLAT, which subsequently triggers proteotoxic stress and cell death [[114]26, [115]47, [116]48]. Another critical hallmark of cuproptosis is the decreased expression of the iron-sulfur cluster protein FDX1, an upstream regulator of protein lipoylation, leading to instability of downstream lipoic acid synthase (LIAS) and consequent mitochondrial dysfunction [[117]25, [118]49] (Fig. [119]4A). Thus, the disruption of intracellular copper homeostasis is a prerequisite for the induction of cuproptosis. The aforementioned experiments have already confirmed that HA-PEG@CuO[2] possessed excellent stability and targeting capabilities, rendering it a promising carrier for the targeted delivery of copper. To determine whether HA-PEG@CuO[2] can induce cuproptosis, we next analyzed the expression of related proteins through WB. As illustrated in Fig. [120]4B, compared to the Control and CuCl[2] group, HA-PEG@CuO[2] significantly induced the oligomerization of DLAT and the instability of FDX1 and LIAS, with their expressions notably downregulated. These findings suggested that HA-PEG@CuO[2] indeed triggered cuproptosis in tumor cells. Fig. 4. [121]Fig. 4 [122]Open in a new tab Effects of HA-PEG@CuO[2] on mitochondrial dysfunction and radiosensitization in vitro. A Schematic illustration of HA-PEG@CuO[2] induced mitochondrial dysfunction by triggering cuproptosis. B Western blot analysis of the key biomarkers of cuproptosis (DLAT, FDX1 and LIAS) in cells after incubated with CuCl[2] and HA-PEG@CuO[2]. C CLSM images of mitochondrial membrane potentials of cells after being incubated with CuCl[2] and HA-PEG@CuO[2] either individually or combined with X-ray. Scale bar = 50 µm. (Red: JC-1 aggregates; Green: JC-1 monomer; Blue: Hoechst). D Bio-TEM images of SMMC 7721 cells after different treatments. Scale bars = 1 μm and 200 nm. E Representative ROS fluorescence images and G corresponding fluorescence intensities of cells treated with above-mentioned formulations. Scale bar = 50 µm. (Blue: Hoechst; Green: ROS). F Representative γ-H[2]AX fluorescence images and J corresponding fluorescence intensities of cells treated with above-mentioned formulations. Scale bar = 50 µm. (Blue: DAPI; Green: γ-H[2]AX). H Representative flow cytometry plots and I quantification of intracellular ROS levels of cells after treated through above-mentioned formulations. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) Mitochondria, as the primary organelles generating ROS and regulating cell death and survival, play a crucial role in radiosensitization [[123]50, [124]51]. Cuproptosis can inhibit the TCA cycle, induce mitochondrial dysfunction, and thereby enhance the efficacy of radiotherapy [[125]52]. Therefore, after demonstrating that HA-PEG@CuO[2] could effectively induce and promote cuproptosis, we further investigated its ability to damage mitochondria and enhance radiosensitization at the cellular level. Firstly, mitochondrial membrane potential (MMP) was assessed using JC-1 as a fluorescent probe. As depicted in Fig. [126]4C, the combination group exhibited the most intense green fluorescence (indicating low MMP) and the weakest red fluorescence (indicating high MMP), indicating pronounced depolarization of the mitochondrial membrane. Moreover, to further investigate the intracellular behavior of HA-PEG@CuO[2], we employed bio-transmission electron microscopy (bio-TEM) to observe the morphological changes in organelles before and after HA-PEG@CuO[2] treatment. As shown in Fig. [127]4D, HA-PEG@CuO[2] treatment resulted in significant alterations, including disruption of mitochondrial and endoplasmic reticulum structures and chromatin anomalies. Notably, the mitochondria exhibited pronounced swelling and vacuolization, increased membrane density, and a marked reduction or complete loss of cristae within the inner mitochondrial membrane. A plethora of research has underscored that inducing ROS generation at tumor sites is a primary mechanism for enhancing the efficacy of radiotherapy [[128]53, [129]54]. Therefore, we evaluated the impact of HA-PEG@CuO[2] on intracellular ROS production using the cell-permeable ROS probe, 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA). As depicted in Fig. [130]4E and G, compared to other groups, the HA-PEG@CuO[2] + X-ray combination treatment group exhibited conspicuous green fluorescence, which is indicative of elevated intracellular ROS levels. Further quantitative analysis via flow cytometry revealed that the level of intracellular ROS in the combination treatment group was approximately 6.4 times higher than that in the radiotherapy group (Fig. [131]4H and I). Afterward, the representative biological marker of DNA double-strand breaks, γ-H[2]AX, was used as an indicator to evaluate the extent of radiation-induced DNA damage. The results were highly consistent with the analysis of ROS generation and cytotoxicity, indicating that the combined treatment of HA-PEG@CuO[2] with X-ray irradiation triggered a heightened level of DNA damage (Fig. [132]4F and J). In summary, our designed HA-PEG@CuO[2] has the potential as a copper-carrier that induces cuproptosis. By triggering cuproptosis and modulating multiple pathways to induce mitochondrial dysfunction, it holds considerable promise for radiosensitization. The transcriptomic and bioinformatics analysis of the mechanism of action of HA-PEG@CuO[2] To investigate the underlying molecular mechanisms of HA-PEG@CuO[2] in enhancing radiosensitization, we conducted RNA-seq analysis on SMMC 7721 cells after different treatments. The mapping rate for each sample ranged from 96.58 to 98.81% (Table S2) and the overall Q30% is above 93.04% (Table S3), suggesting the appropriateness of the reference genome for subsequent bioinformatic analyses. As shown in Fig. [133]5A, the heatmap clearly illustrated the significant overall genetic alterations of tumor cells induced by the different treatment modalities. The volcano plot analysis revealed that radiotherapy resulted in the upregulation of 66 mRNAs and the downregulation of 6 mRNAs (Fig. [134]5B). HA-PEG@CuO[2] treatment led to the upregulation of 2006 mRNAs and the downregulation of 2642 mRNAs (Figure S10A, Supporting Information). When combined with X-ray treatment, HA-PEG@CuO[2] caused the upregulation of 241 mRNAs and downregulation of 1975 mRNAs (Fig. [135]5B). Notably, compared to radiotherapy alone, the combination treatment resulted in the upregulation of 401 mRNAs and the downregulation of 2292 mRNAs, indicating significant changes in gene expression (Figure S10B, Supporting Information). Additionally, a Venn diagram was employed to further analyze the differentially expressed genes present in the X-ray, HA-PEG@CuO[2], and HA-PEG@CuO[2] + X-ray groups (Fig. [136]5C). Fig. 5. [137]Fig. 5 [138]Open in a new tab The transcriptomic and bioinformatics analysis of the mechanism of action of HA-PEG@CuO[2]. A Heatmap of gene expressions in cells after being incubated with CuCl[2] and HA-PEG@CuO[2] individually or in combination with radiotherapy. (1: Control; 2: CuCl[2]; 3: HA-PEG@CuO[2]; 4: X-ray; 5: CuCl[2] + X-ray; 6: HA-PEG@CuO[2] + X-ray). B Volcano plots displayed the differentially expressed genes of X-ray and HA-PEG@CuO[2] + X-ray group compared with Control group. C The differentially expressed genes in Control versus HA-PEG@CuO[2] + X-ray group, Control versus HA-PEG@CuO[2] group, Control versus X-ray group and X-ray versus HA-PEG@CuO[2] + X-ray group in a venn diagram. D KEGG enrichment analysis of the upregulated pathways and E the downregulated pathways of HA-PEG@CuO[2] + X-ray compared to Control group. F GSEA analysis of genes in different pathways. G The O[2] generation curve of HA-PEG@CuO[2] after different treatments at predesigned time points. H Cytotoxicity of HA-PEG@CuO[2] under hypoxia and normoxia. (1: Control; 2: HA-PEG@CuO[2]; 3: X-ray; 4: HA-PEG@CuO[2] + X-ray). I The ATP and J lactate levels in cells after incubated with CuCl[2] and HA-PEG@CuO[2] individually or in combination with radiotherapy. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) To understand the biological functions of the genes that are differentially expressed triggered by the combination treatment, we conducted a pathway enrichment analysis utilizing the Kyoto Encyclopedia of Genes and Genomes (KEGG). Compared to the control group, the combination treatment altered multiple pathways associated with tumor cell proliferation, invasion, metastasis, hypoxia, and metabolism, highlighting its potent anticancer properties (Figure S11, Supporting Information). Firstly, the gene signaling pathways related to cell proliferation and death were significantly affected, including cell cycle signaling pathway, the MAPK signaling pathway, TNF signaling pathway, JAK-STAT signaling pathway and so on. Furthermore, KEGG enrichment analysis illuminated a complex regulatory network of hypoxia-related pathways. Notably, there was a pronounced downregulation of the HIF-1 and NF-kappa B signaling pathways, indicating an improvement of tumor hypoxic microenvironment. This modulation could be attributed to the sustained oxygen release capability of HA-PEG@CuO[2] in response to the tumor microenvironment. Further investigation into the genes expressed revealed their involvement in various metabolic pathways, including TCA cycle, glutathione metabolism, chemical carcinogenesis—reactive oxygen species, nucleotide metabolism, and glycolysis/gluconeogenesis. Such findings underscored the multifaceted impact of the treatment, extending beyond hypoxia amelioration to broad metabolic reprogramming. The PI3K-Akt/mTOR/HIF-1 signaling pathway, a central regulator of glycolysis, plays a critical role in the uptake and utilization of glutamine and glucoses [[139]55]. The tumor suppressor p53 can act as a metabolic regulator by suppressing glycolysis and promoting a shift towards OXPHOS [[140]56]. Specifically, the combination treatment appeared to inhibit glycolysis by inhibiting PI3K/AKT/mTOR/HIF-1α signaling pathway and upregulating p53. Additionally, the upregulation of FoxO was found to have a negative impact on the expression of mitochondrial respiratory chain complexes and OXPHOS, leading to a reduction in cellular ATP content and overall metabolic activity (Fig. [141]5D and E). As expected, Gene Set Enrichment Analysis (GSEA) indicated that chemical carcinogenesis—reactive oxygen species were upregulated, while glutathione metabolism, HIF-1 signaling pathway, glycolysis/gluconeogenesis and related genes were downregulated in the combination treatment group (Fig. [142]5F). These results further elucidated the potential of HA-PEG@CuO[2] in improving tumor hypoxia and mediating tumor metabolism. To further validate the aforementioned transcriptomic sequencing results, we initially utilized a portable dissolved oxygen meter to monitor changes in oxygen concentration over time under different conditions. As illustrated in Fig. [143]5G, the oxygen concentration in the pH 6.8 (10 mM GSH) group was significantly higher than in other groups, indicating the potential of HA-PEG@CuO₂ to produce substantial amounts of oxygen. Comparing the cytotoxicity of HA-PEG@CuO[2] under normoxic and hypoxic conditions revealed that under hypoxic conditions, HA-PEG@CuO[2] reversed hypoxia-induced radioresistance, enhancing cytotoxicity (Fig. [144]5H, Figure S12, Supporting Information). We assessed the expression of HIF-1α using WB after different treatments. As shown in Figure S13, Supporting Information, HA-PEG@CuO[2] under hypoxic conditions effectively alleviated cellular hypoxia, leading to downregulated HIF-1α expression. This is consistent with sequencing results, suggesting the potential of HA-PEG@CuO[2] to alleviate the tumor hypoxic microenvironment. Afterwards, to directly study the impact of HA-PEG@CuO[2] on energy metabolism, we assessed the terminal products of energy metabolism, ATP and lactate. As shown in Fig. [145]5I, J and Figure S14, Supporting Information, the HA-PEG@CuO[2] + X-ray combination treatment group had the lowest levels of ATP and lactate. The T1 enhancement and in vivo biodistribution behavior of HA-PEG@CuO[2] The above results demonstrated that HA-PEG@CuO[2] selectively released Cu(II) at tumor sites. Owing to the unpaired electrons in its outermost orbitals, Cu(II) exhibits paramagnetism and can serve as a T1-weighted MR contrast agent [[146]57, [147]58]. We evaluated the in vitro imaging capabilities of both PVP@CuO[2] and HA-PEG@CuO[2] (Fig. [148]6A and Figure S15, Supporting Information). Initially, under neutral pH conditions (pH 7.4), there was no significant difference in T1-weighted imaging intensity between PVP@CuO[2] and HA-PEG@CuO[2.] However, under acidic conditions, PVP@CuO[2] released Cu(II), which enhanced the T1-weighted imaging intensity regardless of the presence of GSH. In contrast, HA-PEG@CuO[2] released Cu(II) only under acidic conditions with 10 mM GSH, thereby enhancing the T1-weighted imaging intensity. Therefore, the tumor microenvironment responsiveness of HA-PEG@CuO[2], combined with the paramagnetism of Cu(II), endowed HA-PEG@CuO[2] with potential as the T1-weighted MR contrast agent. Fig. 6. [149]Fig. 6 [150]Open in a new tab The T1 enhancement and in vivo biodistribution behavior of HA-PEG@CuO[2]. A Representative T1-weighted intensity of Water, PVP@CuO[2], and HA-PEG@CuO[2]. B Schematic illustration of MR imaging capability of HA-PEG@CuO[2] for MRI-guided antitumor therapy. C Representative T1-weighted MR images of SMMC 7721 tumor-bearing nude mice after intravenous injection with CuCl[2] and HA-PEG@CuO[2] within 24 h. D In vivo fluorescence images of SMMC 7721 tumor-bearing nude mice after intravenous injection with IHA-PEG@CuO[2] at different time points. E Ex vivo fluorescence images of IHA-PEG@CuO[2] in tumor and major organs including the heart, liver, spleen, lung and kidney at 24 h post-intravenous injection. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) Before evaluating the in vivo therapeutic efficacy of HA-PEG@CuO[2], we investigated its biodistribution performance using the tumor-bearing nude mice model (Fig. [151]6B). We subsequently monitored the changes in T1 intensity within the tumor tissue 24 h after tail vein injection of CuCl[2] and HA-PEG@CuO[2] to evaluate their targeted delivery capabilities. The T1-weighted imaging intensity in the HA-PEG@CuO[2] group began to increase 2 h post-injection, with a significant enhancement observed at 4 h (Fig. [152]6C and Figure S16, Supporting Information). This indicated a sustained increase in Cu(II) concentration within the tumor tissue, resulting from the passive targeting properties of nanomaterials, known as the enhanced permeability and retention (EPR) effect and the active targeting effect of HA modification. Conversely, following CuCl[2] injection, there was minimal alteration in the T1-weighted signal intensity at the tumor site. Distinguished by its high temporal resolution and sensitivity, fluorescence imaging serves as an effective tool for investigating the dynamic distribution of nanoparticles in vivo [[153]59, [154]60]. Thus, we employed fluorescence imaging to further evaluate the in vivo biodistrbution performance of HA-PEG@CuO[2]. After intravenous injection of IHA-PEG@CuO[2], imaging was conducted at various time points. As anticipated, a significant ICG fluorescent signal was observed in the tumor tissue over time, which persisted for an extended period (Fig. [155]6D). At 24 h, the mice were euthanized, and the major organs along with tumor tissues were collected for ex vivo fluorescence imaging (Fig. [156]6E), revealing the strongest fluorescence signal in tumor tissues, markedly higher than those in the liver and kidneys. In summary, the dual-modal imaging results from MR and fluorescence imaging demonstrated that HA-PEG@CuO[2] could selectively and rapidly accumulate in the tumor site and responsively release Cu(II), thereby enhancing MR signal. This capability facilitated real-time monitoring of drug delivery to the tumor site. Synergistic antitumor effects of HA-PEG@CuO[2] in combination with LDRT for in situ HCC mouse model Encouraged by the remarkable in vitro anti-tumor effects and substantial tumor accumulation of HA-PEG@CuO[2], we further evaluated its in vivo radiosensitization and anti-tumor efficacy. To closely mimic human hepatocellular carcinoma and assess therapeutic effectiveness, we established the in situ HCC mouse model within the original liver organ. Specifically, SMMC 7721 cells were transplanted into the liver tissue of nude mice to construct the in situ HCC mouse model. Biochemical analyses revealed a notable elevation in the AST and ALT levels within the modeling group (Figure S17A, Supporting Information). Following euthanasia, histopathological examination of murine liver tissues via H&E staining further validated the successful construction of the in situ HCC mouse model (Figure S17B, Supporting Information). Upon successful model establishment, the mice were randomly divided into six groups: Control (PBS), CuCl[2], and HA-PEG@CuO[2] groups, with or without RT. Treatment protocols are shown in Fig. [157]7A. On days 1, 6 and 11, mice were administered the respective nanomaterials (4 mg/kg), followed by radiotherapy (2 Gy) on days 2, 7 and 12. Mice body weights were monitored every day throughout the duration of the experiment. Concurrently, tumor growth was monitored on days 0, 5, 10, 15 and 20 using MR and fluorescence imaging to provide visual representation of tumor volume changes throughout the treatment period. Finally, upon euthanasia, tumors and liver tissues were excised for further analysis. Fig. 7. [158]Fig. 7 [159]Open in a new tab Synergistic Antitumor Effects of HA-PEG@CuO[2] and LDRT for In Situ HCC Mouse Model. A Treatment schedule for the investigation of HA-PEG@CuO[2]-mediated activation of cuproptosis and antitumor effects enhanced RT. B Bioluminescence images and C intensity of in situ HCC mouse model collected at days 0, 5, 10, 15 and 20 after different treatments. D Representative T1-weighted MR images of in situ HCC mouse model and F relative tumor volume changes of mice performed by 3.0 T MR during different treatments. E Representative images of tumors after different treatments. G Average body weight following different treatments. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) As illustrated in Fig. [160]7B–D, over a span of 20 days, the tumor volume in the control group exhibited rapid proliferation, while moderate growth inhibition was observed in HA-PEG@CuO[2], X-ray and CuCl[2] + X-ray groups. In comparison, the tumor growth in the HA-PEG@CuO[2] + X-ray group was significantly suppressed, with a final relative tumor volume of 0.5-fold, which was significantly lower than that of the control group (94.3-fold), CuCl[2] group (93.4-fold), HA-PEG@CuO[2] group (18.3-fold), X-ray group (40.4-fold), and CuCl[2] + X-ray group (40.9-fold) (Fig. [161]7F). Gross specimens of tumors and liver tissues obtained post-euthanasia of mice further corroborated the synergistic antitumor effects of HA-PEG@CuO[2] + X-ray combination therapy, demonstrating superior tumor growth inhibition (Fig. [162]7E). In vitro studies have demonstrated that HA-PEG@CuO[2] exhibited an exceptional ability to release oxygen. Therefore, we employed MR to assess blood oxygenation levels within the tumor regions, aiming to determine whether HA-PEG@CuO[2] could alleviate the hypoxic conditions typical of solid tumors in vivo. Our findings indicated that the T2* values, which reflect tumor oxygenation, were significantly higher in the HA-PEG@CuO[2] group compared to the control and CuCl[2] groups, indicating enhanced tissue oxygenation (Fig. [163]8A and B). Furthermore, WB demonstrated a substantial decrease in HIF-1α expression in the HA-PEG@CuO[2] group, implying significant therapeutic efficacy in alleviating the hypoxic microenvironment of solid tumors (Fig. [164]8C). Fig. 8. [165]Fig. 8 [166]Open in a new tab The mechanisms of synergistic antitumor effects of HA-PEG@CuO[2] and radiotherapy in vivo. A Representative T2* MR images and B T2* values of tumor region of SMMC 7721 tumor-bearing nude mice after different treatments. C Western blot analysis of HIF-1α in tumor cells from SMMC 7721 tumor-bearing mice after different treatments. D Western blot analysis of key biomarkers of the cuproptosis (DLAT, FDX1 and LIAS) in tumor cells from SMMC 7721 tumor-bearing nude mice after different treatments. E Schematic illustration of the process for analyzing tumor tissue sections. F Corresponding fluorescence intensities of DLAT, FDX1 and LIAS of tumor tissues collected from SMMC 7721 tumor-bearing nude mice after different treatments. G Representative fluorescence images and H corresponding fluorescence intensities of H&E staining, ROS, HIF-1α, and γ-H[2]AX of tumor tissues. Scale bar = 50 µm. Data are shown as mean ± SD (n = 3). Statistical analysis was conducted using one-way ANOVA (ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001) To gain deeper insights into the antitumor mechanisms and efficacy of HA-PEG@CuO[2] + X-ray group, we collected tumor tissues for WB, hematoxylin and eosin (H&E), and immunofluorescence analysis (Fig. [167]8E). Firstly, WB analysis of key proteins involved in cuproptosis, including DLAT, FDX1 and LIAS, revealed pronounced aggregation of DLAT and a loss of FDX1 and LIAS in HA-PEG@CuO[2]-treated tumors (Fig. [168]8D). Concurrently, fluorescence staining of relevant proteins further confirmed the occurrence of cuproptosis in vivo (Fig. [169]8F and Figure S18, Supporting Information). Subsequently, H&E staining and immunofluorescence analysis of ROS, HIF-1α, and γ-H[2]AX were performed (Fig. [170]8G, H). As anticipated, H&E staining revealed significant cellular shrinkage and nuclear condensation in the HA-PEG@CuO[2] + X-ray group, along with a marked reduction in tumor cell numbers compared to other groups. As anticipated, this group also exhibited elevated ROS levels, indicating increased oxidative stress. Furthermore, immunofluorescence images of HIF-1α in tumor tissues revealed a significant improvement in tumor hypoxia microenvironment. Finally, γ-H[2]AX staining showed that the green fluorescence intensity in the HA-PEG@CuO[2] + X-ray group was significantly higher than that in the other groups, further confirming its superior antitumor efficacy. In summary, the collective evidence demonstrated that HA-PEG@CuO[2] + X-ray combination therapy significantly ameliorated the hypoxic microenvironment of tumor. This combination strategy elicited a synergistic effect by disrupting mitochondrial function, activating cuproptosis and enhancing radiosensitivity. Consequently, it induced substantial DNA damage, ultimately inhibiting cell proliferation and tumor growth. In vivo biocompatibility and toxicity studies of HA-PEG@CuO[2] Assessing the systemic toxicity of nanomaterials is paramount for determining their viability in clinical applications [[171]61–[172]63]. Thus, we comprehensively assessed the in vivo drug toxicity of HA-PEG@CuO[2] to ensure their safety as potential radiosensitizers. Initially, the mice body weights were monitored throughout the entire administration period, revealing no significant weight loss across all groups (Fig. [173]7G), indicating low toxicity and good biocompatibility. Subsequent histological and biochemical analyses of major organ tissues and blood parameters were conducted to evaluate their potential biological toxicity. Histological staining images indicated no significant abnormalities in the heart, liver, spleen, lung, and kidney among all treatment groups (Figure S19, Supporting Information). Blood biochemical parameters including WBC, HCT, MCV, HGB, RBC, PLT, ALT, AST and CR exhibited no significant differences among all treatment groups (Figure S20, Supporting Information). In conclusion, these results collectively affirmed the reassuring biosafety of HA-PEG@CuO[2] for sensitizing to radiotherapy and facilitating their application in long-term cancer therapy. Discussion Radiotherapy, a mainstream treatment for tumors, has long been limited by radiotherapy resistance and the damage inflicted on surrounding healthy tissues by high-dose X-rays [[174]64]. In recent years, increasing attention has been focused on LDRT [[175]8, [176]65]. Here, we introduced a novel low-dose radiotherapy sensitization approach that improved therapeutic outcomes by inducing cuproptosis and modulating tumor metabolism, with particular emphasis on its application in in situ HCC mouse model. An increasing number of studies suggest that targeting cuproptosis represents a promising radiosensitization strategy to enhance therapeutic efficacy [[177]66–[178]68]. Liao et al. discovered that key regulatory factors of cuproptosis, such as FDX1 and LIAS, are upregulated in residual tumors following radiotherapy, thereby increasing sensitivity to cuproptosis [[179]67]. Moreover, Li et al. demonstrated that a multimodal nanotherapeutic platform promoting cuproptosis could effectively enhance the radiotherapy-induced immunogenic cell death (ICD) effect, achieving over 90% tumor growth inhibition [[180]68]. In our study, both in vitro and in vivo experiments revealed that HA-PEG@CuO[2] effectively triggered cuproptosis and improved therapeutic efficacy. Mitochondria, as essential cellular organelles, are pivotal in energy regulation and serve as convergence points for numerous lethal signal transduction pathways. Our work emphasized the critical role of mitochondria in radiotherapy [[181]17, [182]69–[183]71]. It has been reported that cells dependent on mitochondrial respiration exhibit nearly 1,000 times greater sensitivity to cuproptosis compared to cells reliant on glycolysis [[184]52, [185]72]. Our experiments have confirmed that HA-PEG@CuO[2] continuously released oxygen and copper at the tumor sites, effectively alleviating hypoxia and shifting metabolism from glycolysis to OXPHOS, thus enhancing the sensitivity of tumor cells to cuproptosis. Interestingly, cuproptosis, in turn, caused mitochondrial dysfunction and disrupted OXPHOS, which inhibited energy metabolism and tumor cell proliferation. In this context, we further investigated the structural and functional changes in mitochondria following HA-PEG@CuO[2] treatment, characterized by low membrane potential and elevated ROS levels. The generation of ROS at the tumor sites is a key mechanism for evaluating radiosensitization. Our in vitro experiments showed that ROS levels were significantly higher in the combination therapy group compared to radiotherapy alone. This increase can be attributed to two mechanisms: (1) HA-PEG@CuO[2]-mediated Fenton-like reaction exacerbated oxidative stress and GSH depletion, which increased intracellular ROS production; (2) HA-PEG@CuO[2] activated cuproptosis, leading to mitochondrial dysfunction and aberrant ROS metabolism within tumor cells. Given the increasing clinical demands, innovative biomedical technologies that integrate both disease diagnosis and treatment have gradually emerged as a dominant trend [[186]73–[187]75]. Our study achieved dual-modal fluorescence and MR imaging. Both in vitro and in vivo experiments demonstrated that this approach offered higher sensitivity and resolution compared to conventional MR imaging. This capability facilitated the precise tumor targeting and real-time evaluation of the biodistribution of the nanosystem in vivo. Although HA-PEG@CuO[2] has achieved promising results, it does face potential limitations and presents areas for future investigation. First, while the antitumor efficacy has been demonstrated, additional in vivo studies are required to facilitate its translation into clinical applications. Second, the limited number of experimental animals and the use of immunodeficient mice present constraints, as the interactions between HA-PEG@CuO[2] and the broader tumor-immune microenvironment were not fully explored. Third, future research should investigate the roles of cuproptosis, oxidative stress, and energy metabolism across various cancer types and other potential diseases in clinical patients. Identifying relevant biomarkers to optimize therapeutic strategies is also critical. Additionally, the potential synergistic effects of cuproptosis with radiotherapy and immunotherapy warrant further exploration. Conclusion In summary, we have developed an innovative theranostic nanoplatform (HA-PEG@CuO[2]), which not only modulated tumor metabolism and enhanced cuproptosis sensitivity through the dual enhancement of hypoxia reversal and GSH depletion, but also effectively induced radiosensitization leading to a robust antitumor effect. In vitro experiments demonstrated that HA-PEG@CuO[2], with excellent targeting and tumor microenvironment responsiveness, could mediate the Fenton-like reaction to continuously supply oxygen and Cu(II) while depleting GSH. This process not only induced cuproptosis but also resulted in mitochondrial damage, excessive ROS generation, and inhibition of energy supply, further promoting cell death. Subsequently, bioinformatics analysis confirmed the potential antitumor mechanisms of HA-PEG@CuO[2]. Further, the in situ HCC mouse model was established to evaluate the in vivo antitumor efficacy. Post intravenous injection, MR and fluorescence imaging confirmed that HA-PEG@CuO₂ effectively accumulated and controllably released in the tumor region, demonstrating its dual-modal imaging capability. Both in vitro and in vivo experiments consistently showed that HA-PEG@CuO₂ possessed excellent antitumor effects without noticeable systemic toxicity. In conclusion, this innovative cuproptosis combined with LDRT offered a promising new strategy to overcome radioresistance and improve therapeutic outcomes in hypoxic tumors, particularly HCC. Supplementary Information [188]Additional file 1.^ (10.9MB, docx) [189]Additional file 2.^ (81.5KB, pdf) [190]Additional file 3.^ (64.5KB, pdf) [191]Additional file 4.^ (88.3KB, pdf) Abbreviations bio-TEM Bio-transmission electron microscopy CD44 Cluster of differentiation 44 CLSM Confocal laser scanning microscope DCFH-DA 2',7'-Dichlorodihydrofluorescein diacetate DLAT Dihydrolipoamide S-acetyltransferase DLS Dynamic light scattering DTNB 5,5′-Dithiobis-(2-nitrobenzoic acid) EPR Enhanced permeability and retention effect FDX1 Ferredoxin FTIR Fourier-transform infrared GSEA Gene set enrichment analysis H[2]O[2] Hydrogen peroxide HA Hyaluronic acid HCC Hepatocellular carcinoma KEGG Kyoto encyclopedia of genes and genomes LDRT Low-dose radiation therapy LIAS Lipoic acid synthase MB Methylene blue MMP Mitochondrial membrane potential OXPHOS Oxidative phosphorylation PVP Polyvinylpyrrolidone RBCs Red blood cells RT Radiotherapy TEM Transmission electron microscope TMEs Tumor microenvironments TNB 2-Nitro-5-thiobenzoic acid XPS X-ray photoelectron spectroscopy Author contributions N.S., Y.Q.Y. and G.W.H. contributed equally to this work. H.Z., L.P.L. and Z.Y.X. conceived this research. N.S. and Y.Y.H. synthesized the nanoparticles, conducted the preparation reaction and performed structure characterization. Y.Q.Y., G.W.H and Q.L. directed biological experiments and compiled the figures. N.S., N.L.C. and J.F.C. carried out the cell experiments. N.S. helped to perform animal experiments. N.S., Y.Q.Y., and G.W.H. prepared the manuscript. All of the authors read and approved the manuscript. Funding This work was financially supported by the Guangdong Basic and Applied Basic Research Foundation (2023A1515012660, 2022A1515220159 and 2023A1515220128), the Medical Joint Fund of Jinan University (YXJC2022008), the Guangzhou Science and Technology Plan Project (2023A03J1037), the Clinical Frontier Technology Program of the First Affiliated Hospital of Jinan University, China (JNU1AF-CFTP-2022-a01233) and the National Natural Science Foundation of China (82271943 and 82171893). Availability of data and materials The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. Declarations Ethics approval and consent to participate All animal experiments were conducted in accordance with procedures approved by the Laboratory Animal Welfare and Ethics Committee of Jinan University (IACUC- 20240104–06). Figures were created with BioRender.com. Consent for publication Not applicable. Competing interests The authors declare that they have no competing interests. Footnotes Publisher's Note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Ni Shao, Yongqing Yang and Genwen Hu have contributed equally to this work. Contributor Information Hong Zhang, Email: tzhhong@jnu.edu.cn. Liangping Luo, Email: tluolp@jnu.edu.cn. Zeyu Xiao, Email: zeyuxiao@jnu.edu.cn. References