Introduction
As an indispensable micronutrient, copper exerts vital biological functions in maintaining cellular energy metabolism and redox homeostasis. Disruption of copper homeostasis can impair tumor cell growth and ultimately induce cell death [1], [2]. Therefore, copper-based therapies that disrupt intracellular copper homeostasis to induce copper overload, thereby interfering with key metabolic processes and triggering cell death, have consequently turned into a prominent focus in cancer immunotherapy [3], [4].
Nevertheless, strategies designed to elevate intracellular copper levels for cancer immunotherapy remain hindered by several bottlenecks. First, although several copper ionophores like disulfiram, elesclomol, and 8-hydroxyquinoline have been designed, their clinical application remains hampered by several limitations. These include a short plasma half-life, rapid systemic clearance, and low in vivo stability, which lead to inefficient intracellular copper delivery.
Furthermore, this inefficiency is compounded by copper efflux mechanisms within cells, further depleting the intracellular copper concentration and ultimately compromising the therapeutic efficacy [5], [6]. Second, the tumor microenvironment (TME) typically contains high glutathione (GSH) levels, creating a potent antioxidant barrier. Excess GSH can chelate copper ions and scavenge reactive oxygen species (ROS) generated by Fenton-like reactions, thereby attenuating copper-mediated cytotoxicity [7], [8].
Third, the lack of a stimuli-responsive release mechanism in small-molecule copper ionophores and some copper-based nanomaterials results in poor tumor specificity. This indiscriminate biodistribution across both malignant and healthy tissues inevitably provokes severe systemic side effects [9], [10]. Consequently, innovative nanotechnology strategies are required to significantly elevate intratumoral copper levels, effectively disrupt the GSH antioxidant system, and achieve stimuli-triggered precise drug release at the tumor site, thereby accomplishing efficient eradication of tumor cells.
A marked accumulation of intracellular copper ions can ultimately lead to malignant cell death by triggering multiple mechanisms. Cuproptosis, an emerging type of regulated cell death, is initiated by copper accumulation within cells, which disrupts homeostasis of mitochondria and triggers protein toxicity stress. Furthermore, this pathway functions as an inducer of immunogenic cell death (ICD), promoting the release of damage-associated molecular patterns (DAMPs) to initiate a potent antitumor immune response [11], [12], [13].
The accumulation of copper ions can also amplify oxidative stress via a Fenton-like reaction, thereby boosting intracellular ROS to further elicit pyroptosis. As an inflammatory variant of regulated cell death, pyroptosis is specifically driven by gasdermin (GSDM) family proteins. A key feature of this process is the formation of pores in the plasma membrane, which allows the release of pro-inflammatory cytokines and triggers strong immune activation [14], [15], [16].
The synergistic activation of cuproptosis and pyroptosis reprograms the immunosuppressive "cold" TME into immune-activated "hot" TME, thereby exerting a potent immunotherapeutic effect [17], [18], [19], [20]. Beyond direct TME remodeling, copper-based therapeutics are frequently accompanied by the upregulation of programmed death-ligand 1 (PD-L1) on tumor cells. While elevated PD-L1 typically facilitates immune evasion, it simultaneously provides a highly targetable vulnerability for anti-PD-L1 (αPD-L1) antibodies [21], [22], [23].
Consequently, integrating copper-driven TME reprogramming with αPD-L1 blockade presents a highly rational combination strategy to overcome monotherapy resistance and maximize the durable efficacy of cancer immunotherapy.
Based on these findings, we developed an innovative copper homeostasis nanomodulator, designated CuTF-DC_AC50. This nanomodulator was synthesized in one step by combining Cu2+, 3,3′-dithiobis(propionylhydrazine) (TPH), and DC_AC50 (an inhibitor of the copper chaperone protein ATOX1), then coated with Pluronic F127 to improve stability and biocompatibility (Scheme 1A). DC_AC50 inhibits ATOX1, blocking Cu+ transfer to copper efflux pumps and thereby promoting intracellular free Cu+ accumulation [24].
TPH acts as a dual-responsive unit: its disulfide bond cleaves under high GSH conditions while consuming GSH, and its hydrazide group becomes protonated in the acidic TME, conferring responsiveness to both high GSH and low pH [25]. The CuTF-DC_AC50 features a "threefold dual" strategy, specifically: (1) a "dual-channel" strategy to synergistically elevate intracellular copper levels; (2) dual GSH depletion (via TPH disulfide cleavage and Cu2+ reduction) to disrupt the antioxidant barrier; (3) dual responsiveness to the high GSH and acidic pH of the TME for precise tumor-specific drug release. Specifically, under the dual stimulation of GSH and acidic pH, the nanomodulator is activated to release Cu2+ and DC_AC50.
Cu2+ is converted into the more reactive Cu+, while DC_AC50 prevents cellular Cu+ efflux (Scheme 1B). Together, they lead to massive intracellular Cu+ accumulation. This Cu+ accumulation triggers cuproptosis and, via Fenton-like reactions, elevates ROS levels to induce pyroptosis (Scheme 1C).
The combined cuproptosis/pyroptosis effect robustly induces ICD, releasing DAMPs that promote dendritic cell (DC) maturation and subsequent CD8+ T cell infiltration, reverse the immunosuppressive TME dominated by Tregs and M2-type macrophages, thereby transforming "cold" tumors to "hot" tumors. Given the upregulation of PD-L1 on cancer cells during treatment, we combined this nanomodulator with αPD-L1 therapy in a CT26 tumor model, yielding a collaborative improvement of antitumor immune efficacy (Scheme 1D). In summary, the CuTF-DC_AC50 nanomodulator mediates a targeted copper surge via a "threefold dual" strategy, thereby effectively inducing tumor cell death and stimulating a robust immune response.
By bridging copper-induced sensitization with αPD-L1 blockade, this study presents a robust approach to advance cancer immunotherapy.Scheme 1Schematic depiction of a copper homeostasis nanomodulator to trigger cuproptosis/pyroptosis and its combination with αPD-L1 for enhanced antitumor immunity. (A) Scheme of the preparation process of CuTF-DC_AC50 via self-assembly method. (B) Schematic illustration of the "threefold dual" strategy. i.
Dual-responsive degradation; ii. Dual-channel copper overload; iii. Dual GSH depletion.
(C) Cu+-mediated cuproptosis/pyroptosis inducing ICD. (D) PD-L1 blockade enhances cancer immunotherapy.
Results and discussion
Synthesis and characterization of CuTF-DC_AC50
CuT-DC_AC50 was first synthesized through a one-pot self-assembly method (Fig. 1A), followed by surface coating with the amphiphilic polymer F127 to yield the final CuTF-DC_AC50. Transmission electron microscopy (TEM) revealed that the unmodified CuT-DC_AC50 exhibited a uniform spherical morphology (Fig.
1B), whereas the surface-modified CuTF-DC_AC50 displayed a frosted glass-like surface topography (Fig. 1C). The EDS elemental mapping verified the presence of Cu, S, N, O, and F elements, clearly validating the successful fabrication of CuT-DC_AC50 (Fig.
1D). Dynamic light scattering (DLS) analysis demonstrated that CuTF-DC_AC50 possessed a hydrodynamic diameter of approximately 106 nm, coupled with an exceptionally narrow size distribution, as evidenced by a polydispersity index (PDI) of 0.075, demonstrating excellent mono-dispersity beneficial for enhanced penetration and retention (EPR) effect (Fig. 1E).
Zeta potential analysis further confirmed that the CuT-DC_AC50 surface carried a positive charge (21.4 mV), which shifted to near-neutral following F127 modification (0.15 mV) (Fig. 1F). Such surface charge characteristics can reduce non-specific protein adsorption under physiological conditions and prolong systemic circulation time.
To elucidate the element makeup and chemical bonding states within the CuT framework, an X-ray photoelectron spectroscopy (XPS) was conducted. The spectrum verified the existence of Cu, C, N, O, and S (Fig. 1G).
The high-resolution Cu 2p spectrum exhibits two characteristic Cu 2p3/2 peaks at 934.9 eV and 932.8 eV, each accompanied by a satellite peak, indicating that copper exists as Cu2+ in two distinct coordination environments (Fig. 1H). The high-resolution N 1s spectrum was deconvoluted into two peaks at 400.6 eV and 399.3 eV (Fig.
1I), which were assigned to N-H and NH2 of hydrazide groups, respectively. Compared with that of free TPH, the NH2 peak shifts toward lower binding energy by approximately 0.6 eV, indicating coordination between Cu2+ and the terminal N of the hydrazide group [26]. The deconvolution of the S 2p3/2 spectrum revealed two peaks at 163.5 eV and 162.3 eV, corresponding to uncoordinated S and coordinated S, respectively, with a predominance of uncoordinated S species (Fig.
1J). The high-resolution O 1s spectrum shows two distinct peaks at 532.6 eV and 531.0 eV, attributed to C–O and Cu–O bonds, respectively (Fig. S1).
These results demonstrate that CuT is formed via the binding of Cu2+ to N, S and O in TPH. Fourier transform infrared (FTIR) spectroscopy further supports the coordination mode (Fig. S2).
Based on the XPS and FTIR analysis results, the proposed coordination structure of CuT is illustrated in Fig. S3.Fig. 1Synthesis and structural characterization of CuTF-DC_AC50.
(A) Schematic illustration of the synthetic procedure of CuTF-DC_AC50. (B) TEM image of CuT-DC_AC50. Inset: HRTEM image of CuT-DC_AC50.
Scale bar: 10 nm. (C) TEM micrograph of CuTF-DC_AC50. Inset: HRTEM image of CuTF-DC_AC50.
Scale bar: 10 nm. (D) Element mappings of CuT-DC_AC50. (E) Size distribution of CuTF-DC_AC50 determined by DLS.
(F) Zeta potential of CuT-DC_AC50, F127 and CuTF-DC_AC50. (G) Survey XPS spectrum, high-resolution (H) Cu 2p, (I) N 1s and (J) S 2p XPS spectra of CuT. (K) FTIR spectrum of CuT, DC_AC50, CuT-DC_AC50, and CuTF-DC_AC50.
(L) The N2 adsorption–desorption isotherm of CuT. (M) Time-evolved particle size stability of CuTF-DC_AC50 dispersed in H2O, PBS, and RPMI-1640.
FTIR spectroscopy was also employed to verify DC_AC50 incorporation and F127 modification (Fig. 1K). An absorption band at 3308 cm−1 detected in the CuT-DC_AC50 arises from the N–H stretching mode of amide structural groups.
For both CuT-DC_AC50 and CuTF-DC_AC50, the spectral signal near 1600 cm−1 is associated with the skeleton vibrational modes of aromatic C=C chemical bonds. The appearance of these characteristic absorption bands confirms the successful loading of DC_AC50. Furthermore, the characteristic peak observed at 2888 cm−1 for CuTF-DC_AC50 is assigned to the C–H stretching vibration, providing direct evidence for successful F127 surface coating.
Fluorescence spectroscopy analysis revealed that upon excitation at 355 nm, CuTF-DC_AC50 exhibited a distinct emission peak at 472 nm (Fig. S4), further corroborating the successful loading of DC_AC50 [27].
To investigate the drug loading mechanism, the pore structure of the drug-free CuT framework was characterized by nitrogen adsorption-desorption measurements. The adsorption-desorption isotherm displayed a typical hysteresis loop, which indicates a mesoporous architecture within the nanomodulator. Brunauer-Emmett-Teller (BET) analysis showed that the specific surface area, pore volume and pore size were determined to be 78.88 m2 g−1, 0.316 cm3 g−1 and 4.9 nm, respectively (Fig.
1L). The loading efficiency of DC_AC50 was calculated at roughly 5% using a steady-state and transient fluorescence spectrometer combined with a standard curve method (Fig. 2K and Fig.
S5). Additionally, quantitative assessment by inductively coupled plasma mass spectrometry (ICP-MS) indicated that the copper content in CuT-DC_AC50 reached approximately 21%. The stability of CuTF-DC_AC50 was evaluated over 7 d in ultrapure water, phosphate-buffered saline (PBS), and RPMI-1640 cell culture medium (Fig.
1M). The results demonstrated that CuTF-DC_AC50 maintained homogeneous dispersion without noticeable aggregation or precipitation throughout the testing period, confirming its excellent physical stability and providing a reliable foundation for subsequent biological experiments.Fig. 2Physicochemical properties of CuTF-DC_AC50.
(A) Schematic diagram of GSH/pH-dual-responsive degradation of CuTF-DC_AC50. (B) Depletion of GSH detected by DTNB at various concentrations of CuTF-DC_AC50 and (C) different reaction times. (D) ·OH production at different concentrations of CuTF-DC_AC50 by MB.
(E) ·OH production at different reaction times by MB. Inset: the color changes of the solutions. (F) MB degradation monitored by absorbance at 664 nm after incubation with CuTF-DC_AC50, H2O2, and different GSH concentrations at pH 6.5.
(G) Schematic illustration showing (a) dual GSH consumption of CuTF-DC_AC50 (involving disulfide bond cleavage and reduction of Cu2+) and (b) subsequent Cu+-mediated Fenton-like reaction for ROS generation. (H) TEM images of CuTF-DC_AC50 after different treatments for 24 h. Cumulative release of (I) Cu2+ and (J) DC_AC50 from CuTF-DC_AC50 under varying pH conditions with or without GSH.
(K) Concentration-dependent fluorescence spectra of DC_AC50.
Physicochemical properties of CuTF-DC_AC50
The physicochemical properties of CuTF-DC_AC50 were thoroughly characterized, focusing on its dual pH/GSH responsiveness (Fig. 2A), its capacity for GSH depletion and Fenton-like reaction-driven ROS generation. GSH levels are distinctly higher in malignant cells than in normal cells, a biochemical alteration that facilitates tumor progression by preserving intracellular redox equilibrium [28], [29], [30].
5,5′-dithiobis(2-nitrobenzoic acid) (DTNB) was employed to evaluate the GSH-depleting efficacy of CuTF-DC_AC50. According to Fig. 2B and C, the characteristic absorbance of the reaction product TNB at 412 nm decreased significantly with increasing nanoparticle concentration or prolonged incubation, confirming effective GSH consumption and the disruption of tumor cells antioxidant defense by CuTF-DC_AC50.
Methylene blue (MB) served as a molecular indicator for the detection of hydroxyl radical (•OH) generation from Fenton-like reactions triggered by CuTF-DC_AC50. The absorbance of MB at 664 nm exhibited a concentration- and time-dependent decrease, confirming the efficient generation of •OH (Fig. 2D and E).
Furthermore, the influence of GSH concentration on •OH yield was investigated. As illustrated in Fig. 2F, •OH production increased with rising GSH concentrations from 0 to 2 mM but decreased at higher GSH levels, likely owing to radical scavenging by excess GSH—a finding consistent with previous reports [31].
Notably, partial MB degradation was observed even in the absence of GSH, which can be attributed to acidic pH-triggered disassembly of the nanomodulator and the subsequent initiation of a Fenton-like reaction. Collectively, these results demonstrate that CuTF-DC_AC50 effectively depletes GSH and generates toxic •OH through a Fenton-like reaction (Fig. 2G).
The stimulus-responsive degradation and release profile of CuTF-DC_AC50 were further investigated. TEM images (Fig. 2H) visually confirmed morphological changes after 24 h incubation upon different treatments.
At physiological neutral pH (7.4), the nanomodulator maintained a spherical morphology, whereas obvious structural disintegration was observed under simulated TME conditions, including acidic pH (6.5) or high GSH levels (pH 7.4 with GSH). The complete degradation of the nanomodulator occurred in response to acidic pH and high GSH dual stimuli (pH 6.5 + GSH), revealing their synergistic responsiveness to TME acidity and elevated GSH. The quantitative release profiles of Cu2+ and DC_AC50 were investigated (Fig.
2I and J). Only minimal release of Cu2+ (∼3%) and DC_AC50 (∼5%) was detected at physiological neutral pH (7.4) over 24 h, suggesting excellent stability under physiological conditions. In contrast, drug release was significantly accelerated under single-stimulus conditions (pH 6.5 or GSH).
Under weakly acidic conditions (pH 6.5 alone), 13% of Cu2+ and 23% of DC_AC50 were released within 24 h, whereas under GSH-only conditions, 42% of Cu2+ and 54% of DC_AC50 were released within the same time period. Upon exposure to dual stimuli (pH 6.5 + GSH), the cumulative release rates reached ∼69% for Cu2+ and ∼85% for DC_AC50, further confirming the dual-responsive behavior of the nanomodulator in the TME. In summary, CuTF-DC_AC50 exhibits intelligent TME dual-responsive behavior, efficient GSH depletion, and potent •OH-generating capability—laying a reliable physicochemical basis for further antitumor research.
Evaluation of cellular internalization and antitumor efficacy in vitro
Given its dual responsiveness to the TME (acidic pH/high GSH) and its ability to produce •OH radicals through Fenton-like reactions, the cellular internalization of FITC-labeled CuTF-DC_AC50 by CT26 cells was first evaluated using confocal laser scanning microscopy (CLSM). The time-dependent enhancement of green fluorescence signals, as evidenced in Fig. 3A and Fig.
S6, indicates that the nanomodulator was successfully internalized by the cells. Efficient lysosomal escape is essential for preventing the degradation of nanoparticles within lysosomes and ensuring their effective delivery to the cytoplasm, where their therapeutic action occurs [32], [33], [34], [35]. To assess whether CuTF-DC_AC50 could successfully escape from lysosomes following cellular internalization, colocalization between the lysosomal fluorescent probe LysoTracker Red and FITC-labeled nanoparticles was first examined.
The results revealed that after 4 h of incubation, partial accumulation of the nanomodulator within lysosomes was observed. After 6 h of incubation, significant colocalization was detected between the nanomodulator and lysosomes, with a Pearson's correlation coefficient of 0.73, confirming that a substantial portion of the nanoparticles remained enclosed within lysosomes. Notably, after 8 h of incubation, a reduction in the LysoTracker Red signal and diminished colocalization were observed, indicating lysosomal dysfunction that facilitates the release of nanoparticles into the cytoplasm (Fig.
3C). Bio-transmission electron microscopy (Bio-TEM) was subsequently employed to further corroborate the occurrence of lysosomal escape. As shown in Fig.
S7, after 4 h of co-incubation, CuTF-DC_AC50 exhibited partial aggregation within lysosomes. After 6 h of incubation, the nanoparticles remained localized within lysosomes, although some lysosomal membranes displayed blurred contours. After 8 h of incubation, clusters of nanoparticles were observed dispersed in the cytoplasm, demonstrating successful lysosomal escape.Fig.
3Cellular uptake and cytotoxicity. (A) Cellular uptake images of CuTF-DC_AC50 incubated with CT26 cells at various time intervals. (B) Survival rates of CT26 and NIH 3T3 cells upon 24 h treatment with rising concentrations of CuTF-DC_AC50.
(C) Analysis of co-localization between FITC-labeled CuTF-DC_AC50 (green) and lysosomal markers (red) in CT26 cells. (D) Survival rates of CT26 cells upon 24 h incubation with CuTF-DC_AC50 under diverse conditions. (E) Survival rates of CT26 cells upon 24 h incubation with distinct formulations.
(F) The live & dead staining and fluorescence statistics of CT26 cells with distinct formulations. (G) Apoptotic analysis via flow cytometry in Annexin V-FITC/PI double-labeled CT26 cells after various treatments. (H) Relative GSH content in CT26 cells after various treatments.
(I) Fluorescence imaging and quantitative analysis of ROS production in CT26 cells following various treatments. Data are shown as mean ± SD (n = 3). p values were derived from a one-way ANOVA test, **p < 0.01, ***p < 0.001, ****p < 0.0001.
The in vitro antitumor efficacy of CuTF-DC_AC50 was then evaluated. At physiological pH 7.4, the nanomodulator exhibited excellent cytocompatibility, with cell viability remaining above 80% in both CT26 and mouse embryonic fibroblast (NIH/3T3) cells across a range of concentrations (Fig. 3B).
In contrast, when CT26 cells were exposed to conditions mimicking the acidic extracellular TME (pH 6.5) or treated with exogenous 5 mM GSH as a substrate to verify GSH responsiveness (pH 7.4 + 5 mM GSH), cell viability decreased in a concentration-dependent manner (Fig. 3D). The most pronounced cytotoxic effect was observed under combined stimulation (pH 6.5 + 5 mM GSH), where cell viability dropped to 9% at 200 μg/mL, highlighting the synergistic responsiveness of the nanomodulator.
Comparative evaluation of different formulations (free DC_AC50, CuTF, and CuTF-DC_AC50) under the same dual-stimulus condition confirmed that CuTF-DC_AC50 exhibited superior concentration-dependent cytotoxicity (Fig. 3E). Live/dead cell staining further verified the strengthened antitumor effect (Fig.
3F), which revealed approximately 80% cell death in the CuTF-DC_AC50 group, markedly higher than those in DC_AC50 (11%) and CuTF (20%) groups. Consistent with these findings, the CuTF-DC_AC50 group also demonstrated the lowest colony-forming ability (Fig. S8).
Apoptosis analysis via Annexin V-FITC/PI staining (Fig. 3G and Fig. S9) revealed that CuTF-DC_AC50 induced 79.0% apoptosis, which was 2.9-fold and 1.3-fold increase against that induced by DC_AC50 (27.1%) and CuTF (61.9%), respectively, highlighting its enhanced capacity to trigger programmed cell death.
High concentrations of GSH (2–10 mM) within tumor cells promote antioxidant defense, immune evasion, and therapeutic resistance [36], [37]. Therefore, depleting intracellular GSH represents a crucial therapeutic strategy. In the CuTF-DC_AC50 group, the disulfide bonds in the TPH moiety and the released Cu2+ act synergistically to deplete GSH efficiently.
Intracellular GSH measurement confirmed that the CuTF-DC_AC50 treatment induced the most significant GSH reduction, with residual GSH levels of 75.5%, 56.3%, and 40.3% in the DC_AC50, CuTF, and CuTF-DC_AC50 groups, respectively (Fig. 3H). The disruption of redox homeostasis resulted in a significant increase in ROS, as tracked by DCFH-DA fluorescence probe.
This increase was most pronounced in the CuTF-DC_AC50-treated group, as verified by prominent green fluorescence intensity (Fig. 3I), confirming that this group had the highest level of ROS accumulation. Further evidence from flow cytometry validated this tendency (Fig.
S10). In addition to investigating the antitumor efficacy across different treatment groups, comparative analyses of CuTF-DC_AC50 under single stimuli (either pH 6.5 or 5 mM GSH alone) were also conducted (Fig. S11–S14).
The results demonstrated that, compared with single-stimulus conditions, CuTF-DC_AC50 exhibited the strongest cytotoxic effect under the dual-stimulus condition (pH 6.5 + 5 mM GSH), mediating the highest levels of GSH depletion and ROS generation.
Oxidative stress triggered severe mitochondrial dysfunction. The JC-1 assay demonstrated a profound loss of mitochondrial membrane potential (ΔΨm) in the CuTF-DC_AC50 group, as indicated by markedly elevated green fluorescence compared with other groups (Fig. 4A and Fig.
S15). Furthermore, MitoTracker Red CMXRos staining showed that CuTF-DC_AC50 treatment caused mitochondrial fragmentation from an elongated tubular morphology to punctate structures (Fig. 4C).
Bio-TEM provided direct ultrastructural evidence of these pathological changes, which included marked mitochondrial swelling, vacuolization, and eventual cell membrane rupture (Fig. 4B). In summary, CuTF-DC_AC50 demonstrates efficient cellular uptake, potent stimulus-dependent cytotoxicity, significant GSH depletion, ROS amplification, and induction of mitochondrial-mediated cell death, establishing it as a promising antitumor candidate in vitro.Fig.
4Antitumor efficacy in vitro. (A) Fluorescence imaging of mitochondrial membrane potential in CT26 cells using JC-1 staining following various treatments. (B) Representative Bio-TEM micrographs illustrating ultrastructural changes in treated CT26 cells.
(C) CLSM images of mitochondria stained with MitoTracker in CT26 cells following the various treatments. Scale bar: 10 μm (overview) and 5 μm (zoom). (D) The "Dual-Channel" strategy (a) and subsequent activation of cuproptosis and pyroptosis pathways (b).
(E) Immunofluorescence analysis depicting FDX1 expression following the various treatments. (F) Immunofluorescence analysis depicting DLAT expression following the various treatments. (G) Protein expression levels of ATOX1, DLAT, FDX1, and PD-L1 assessed by Western blot.
Cuproptosis and pyroptosis converge to trigger ICD and drive DC maturation
The molecular mechanisms underlying the antitumor effects of CuTF-DC_AC50 were subsequently investigated, with an initial focus on its ability to induce cuproptosis. Given that elevated intracellular copper concentration is a decisive trigger for cuproptosis [38], [39], [40], CuTF-DC_AC50 was designed to amplify intracellular copper levels primarily through a "dual-channel" strategy that simultaneously enhances copper influx and inhibits its efflux (Fig. 4D).
Specifically, following its internalization by tumor cells, CuTF-DC_AC50 underwent responsive intracellular degradation. The released Cu2+ serves as an exogenous source to elevate the intracellular copper pool, while the co-released inhibitor DC_AC50 blocks the copper chaperone protein ATOX1, thereby inhibiting copper ion efflux. This synergistic effect elevates intracellular copper levels, thereby triggering cuproptosis.
To validate this hypothesis, the effect of CuTF-DC_AC50 on ATOX1 expression was further examined. Western blot analysis demonstrated a marked reduction in ATOX1 expression after CuTF-DC_AC50 intervention, relative to the control (Fig. 4G), validating the effective intracellular delivery and inhibitory function of the DC_AC50 moiety.
The downregulation of ATOX1 is expected to impair its chaperone function for cuprous ions, leading to intracellular copper retention and blocked efflux.
Key molecular events in the cuproptosis pathway were then investigated. Immunofluorescence and Western blot analyses demonstrated that the CuTF-DC_AC50 treatment significantly reduced both the fluorescence intensity and protein level of ferredoxin 1 (FDX1), a critical upstream modulator of cuproptosis, compared with DC_AC50 or CuTF alone (Fig. 4E–G, and Fig.
S16A). These results suggest that CuTF-DC_AC50 disrupts FDX1 stability, potentially via copper-mediated disassembly of its iron-sulfur cluster. Furthermore, aberrant oligomerization of dihydrolipoamide S-acetyltransferase (DLAT) represents a key feature of cuproptosis.
Immunofluorescence analysis revealed distinct DLAT aggregates in tumor cells treated with CuTF-DC_AC50, contrasting with findings in the other groups (Fig. 4F and Fig. S16B).
Western blot analysis further confirmed the most significant DLAT oligomerization in this group (Fig. 4G). Taken together, the findings indicate that CuTF-DC_AC50 triggers cuproptosis by disrupting copper homeostasis and targeting key executers of the cuproptosis pathway.
Previous research has indicated that copper-based nanomaterials can upregulate PD-L1 expression on tumor cells during cancer therapy [21], [23]. To validate this effect, western blot was conducted to assess PD-L1 expression in differently treated CT26 cells. The results exhibited that CuTF-DC_AC50 treatment induced the most significant upregulation of PD-L1 (Fig.
4G). This finding provides a rationale for the subsequent combination of CuTF-DC_AC50 with αPD-L1 ICB therapy. To further elucidate the molecular mechanisms underlying tumor cell PD-L1 upregulation, CT26 cells were treated with varying concentrations of CuTF-DC_AC50 (0, 12.5, 25, 50, and 100 μg/mL).
The low-concentration groups (12.5 and 25 μg/mL) induced copper stress without causing significant cell death, whereas higher concentrations (50 and 100 μg/mL) resulted in varying degrees of cell death. After treatment, PD-L1 expression levels were assessed by Western blot analysis (Fig. S17).
Compared to the control group, PD-L1 expression was mildly upregulated in the low-concentration groups, indicating that direct intracellular copper stress alone can elevate PD-L1 expression, potentially via activation of the JAK-STAT3 signaling pathway [41]. In contrast, PD-L1 expression was further elevated in the higher concentration groups, suggesting that secondary stress associated with cell death also contributes to PD-L1 upregulation, likely mediated through DAMPs and inflammatory signaling cascades [17], [42].
Next, we investigated whether CuTF-DC_AC50 could simultaneously trigger pyroptosis. A key microscopic hallmark of early-stage pyroptosis is the distinctive blebbing of the plasma membrane [20], [43], [44]. In line with this characteristic, morphological analysis of CT26 cells following CuTF-DC_AC50 treatment revealed classic pyroptotic features, including pronounced cellular swelling and the development of large, balloon-shaped vesicles.
Remarkably, no such morphological variations occurred in the control and DC_AC50 cohorts, while they were hardly noticeable in the CuTF group (Fig. 5A). As an upstream event of pyroptosis, activation of the canonical NLRP3 inflammasome begins with the oligomerization of NLRP3 protein, followed by recruitment of ASC and the formation of ASC specks, leading to caspase-1 activation and gasdermin D (GSDMD)-dependent pore formation, ultimately resulting in pyroptotic cell death [45], [46].
Therefore, NLRP3 expression levels were evaluated. Western blot analysis shown in Fig. S18 revealed a significant upregulation of NLRP3 expression in the CuTF-DC_AC50 group compared to control group, indicating NLRP3 priming.
Immunofluorescence staining demonstrated a marked increase in the number of intracellular ASC specks following CuTF-DC_AC50 treatment (Fig. S19). Subsequent Western blot analyses further showed significantly elevated levels of cleaved caspase-1 (c-caspase-1) and the GSDMD-N-terminal fragment (GSDMD-NT in the CuTF-DC_AC50 group (Fig.
5B). Collectively, these results confirm that CuTF-DC_AC50 activates the canonical NLRP3 inflammasome pathway. During pyroptosis, GSDMD-NT undergoes oligomerization to induce pore formation in cell membranes, triggering sustained release of cytoplasmic contents [47], [48], [49].
Consistent with this mechanism, the CuTF-DC_AC50 group showed substantial lactate dehydrogenase (LDH) release, with only 26% retained intracellularly, compared with 83% and 65% in the DC_AC50 and CuTF groups, respectively (Fig. 5C). Furthermore, the concentration of IL-1β reached 127 pg/mL in the CuTF-DC_AC50 group, marking a 2.4-fold elevation compared to the control group and signifying the most substantial release of the pro-inflammatory cytokine (Fig.
5D). Collectively, these morphological, biochemical, and immunological results confirm that CuTF-DC_AC50 is a potent inducer of pyroptosis, primarily through the caspase-1/GSDMD-dependent axis.Fig. 5Antitumor mechanism in vitro.
(A) Assessment of cellular morphology in CT26 cells after varied interventions. (B) Assessment of pyroptosis-associated protein levels via Western blot. (C) Measurement of intracellular LDH levels in CT26 cells post-treatment.
(D) IL-1β levels in mouse serum following different treatments. (E) Schematic illustration of ICD triggered by cuproptosis and pyroptosis (a), thereby stimulating DC maturation (b). (F) Immunofluorescence analysis depicting CRT expression following the various treatments.
(G) Immunofluorescence analysis depicting HMGB1 expression after varied interventions. (H) Determination of intracellular ATP content in CT26 cells. (I) In vitro DC maturation (CD11c+CD80+CD86+) assessed by flow cytometry after co-culture with treated CT26 cells.
Data are shown as mean ± SD (n = 3). p values were calculated via one-way ANOVA test, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
Given the efficacy of CuTF-DC_AC50 in inducing both cuproptosis and pyroptosis, this study further investigated whether the synergistic interplay between these two pathways could trigger robust ICD, thereby potentially eliciting a potent antitumor immune response (Fig. 5E). ICD is defined by the release of DAMPs that stimulate DC maturation and the subsequent infiltration of CD8+ T cells [50], [51].
The potential of CuTF-DC_AC50 to act as an ICD inducer was evaluated by systematically examining the release of key DAMPs (calreticulin (CRT), high mobility group box 1 (HMGB1), and ATP). Under physiological conditions, CRT is localized to the endoplasmic reticulum (ER) lumen. Upon ER stress induction, it translocates to the plasma membrane where it functions as a key "eat-me" cue for phagocytic cells [52].
Immunofluorescence staining revealed a distinct ring-shaped fluorescence of CRT selectively on the surface of CuTF-DC_AC50-treated cells, confirming robust CRT exposure, in stark contrast to the control groups (Fig. 5F and Fig. S20A).
Under physiological homeostasis, HMGB1 predominantly localizes to the nucleus. In the process of ICD, HMGB1 is discharged from the nucleus into the extracellular environment in a passive manner. Accordingly, obvious decrease in nuclear HMGB1 fluorescence was observed in CuTF-DC_AC50-treated group (Fig.
5G and Fig. S20B). Furthermore, ICD is associated with a rapid depletion of intracellular ATP due to mitochondrial dysfunction.
Consistently, CuTF-DC_AC50 treatment sharply lowered intracellular ATP content to only 5% of the initial level (Fig. 5H). These DAMP release demonstrates that CuTF-DC_AC50 is a potent ICD inducer.
The hallmark of ICD is its ability to activate adaptive immunity through dendritic cells (DCs). We employed a transwell co-culture system to assess DC maturation quantitatively. Bone marrow-derived DCs (BMDCs) were co-cultured with CT26 cells receiving varied interventions, and their expression of the co-stimulatory molecules CD80 and CD86 was analyzed by flow cytometry.
Flow cytometric analysis showed that tumor cells treated with CuTF-DC_AC50 induced the highest BMDC maturation rate (approximately 49.5%), approximately 1.6-fold and 1.5-fold higher relative to DC_AC50 and CuTF cohorts (Fig. 5I). These results conclusively demonstrate that CuTF-DC_AC50-induced ICD effectively drives DC activation and maturation at the cellular level, providing a strong rationale for exploring its potential to elicit antitumor immune responses in vivo.
CuTF-DC_AC50 synergizes with αPD-L1 to potentiate antitumor immunity
Prompted by the observation that CuTF-DC_AC50 upregulates PD-L1 expression, we subsequently combined this nanomodulator with αPD-L1 blockade in vivo to maximize the overall efficacy of antitumor immunotherapy. To evaluate the impact of CuTF-DC_AC50 on immune cells within the immunosuppressive TME, CT26 tumor-bearing female BALB/c mice were randomly assigned to five groups and subjected to a 14 d treatment regimen (Fig. S21): G1.
Control; G2. DC_AC50; G3. CuTF; G4.
CuTF-DC_AC50; and G5. CuTF-DC_AC50 + αPD-L1. Following the treatment period, single-cell suspensions were prepared from the spleens, lymph nodes (LNs), and tumors of each mouse for flow cytometry analysis.
Experimental observations indicated a notable upsurge in DC maturation within the CuTF-DC_AC50 group relative to the control. Notably, the combination therapy (CuTF-DC_AC50 + αPD-L1) achieved the highest maturation rates (25.0% in spleen, 16.3% in LNs, and 66.7% in tumors) (Fig. 6A–F, and Fig.
S22). Upon maturation, DCs migrate to secondary lymphoid organs including the spleen and lymph nodes. Within these sites, antigen presentation by DCs facilitates the infiltration of cytotoxic T lymphocytes [53], [54], [55], [56].
As anticipated, the CuTF-DC_AC50 + αPD-L1 group exhibited a marked elevation of CD8+ T cell versus the control, with increases of 1.8-fold in the spleen, 2.9-fold in tumors, and 1.5-fold in LNs (Fig. 6B–G, and Fig. S23).
The enhanced intratumoral infiltration of CD8+ and CD4+ T cell following this combination therapy was further confirmed by immunofluorescence (Fig. 8H).Fig. 6In vivo immune activation.
(A) Quantitative assessment of splenocyte-derived DC maturation. (B) Quantitative assessment of splenocyte-derived CD8+ T cells. (C) Quantitative assessment of tumor-derived M1 macrophages.
(D) Quantitative assessment of tumor-derived M2 macrophages. (E) Quantitative assessment of lymph node-derived Tregs. (F) Quantitative assessment of lymph node-derived DC maturation.
(G) Quantitative assessment of tumor-derived CD8+ T cells. (H) Quantitative assessment of the ratios of M1/M2 macrophages in the tumor cells with varied interventions (I) Comparative radar chart of immune cell subset expression in splenocytes with various treatments. (J-M) Serum contents of TNF-α (J), IL-6 (K), IFN-γ (L) and IL-10 (M) levels quantified via ELISA.
Data are shown as mean ± SD (n = 3). p values were derived from a one-way ANOVA test, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
M2-type macrophages and Tregs are pivotal immunosuppressive populations within the immunosuppressive TME. The polarization status of TAMs and the proportion of Tregs were evaluated among all experimental cohorts. The obtained data indicated that the combination therapy significantly increased the M1/M2 macrophage ratio in all examined tissues (Fig.
6H, Fig. S24 and S25), along with elevated antitumor M1-type macrophages and diminished protumorigenic M2-type macrophages (Figs. 6C, D, and Fig.
S26-29). Moreover, a sharp decline in Treg ratios was detected in the combined treatment group—by 59% (LNs), 78% (spleen), and 70% (tumor) relative to the control (Fig. 6E, Fig.
S30 and S31). Radar chart comprehensively depicts the expression of immune cell subsets in the spleen, LNs and tumor tissue across different experimental groups (Fig. 6I, Fig.
S32 and S33). This alleviation of immunosuppression was also reflected in serum cytokine expression levels. Compared with other groups, the CuTF-DC_AC50 + αPD-L1 combination therapy significantly enhanced the release of the pro-inflammatory mediators TNF-α, IL-6, and IFN-γ, while significantly reducing the level of immunosuppressive interleukin-10 (IL-10) (Fig.
6J–M). The increased IFN-γ level indicates robust CD8+ T cell effector function, and the elevated TNF-α together with reduced IL-10 reflects a favorable immune shift toward activation rather than suppression. Collectively, the CuTF-DC_AC50 not only potently activates DCs and promotes CD8+ T cell infiltration, but also effectively remodels the immunosuppressive TME by repolarizing TAMs and reducing Tregs.
The combination of CuTF-DC_AC50 with αPD-L1 demonstrates a synergistic effect, significantly enhancing in vivo antitumor immunity.
Biodistribution, biological safety and antitumor efficacy in vivo
NIR-II fluorescence imaging was employed to investigate the tissue distribution profile of CuTF-DC_AC50 in mice bearing CT26 tumors. In vivo imaging showed a time-dependent accumulation of CuTF-DC_AC50 in tumor tissues, with peak fluorescence intensity detected 24 h post-administration (Fig. 7A–C and Fig.
S34). Ex vivo imaging analysis of isolated tumor and major organs obtained 24 h post-treatment confirmed significant accumulation in tumor tissue, with the fluorescence intensity ranking second only to that in the liver (Fig. 7B–D).
These findings demonstrate that CuTF-DC_AC50 exhibits excellent tumor-enriching capacity, characterized by effective accumulation and prolonged retention in tumor tissue. Such a favorable biodistribution profile justified further systematic evaluation of its in vivo antitumor performance.Fig. 7In vivo biodistribution and biosafety.
(A) Non-invasive fluorescence imaging at serial time points post intravenous administration of IR820-labeled CuTF-DC_AC50 in mice bearing CT26 tumors. (B) Representative ex vivo fluorescence images of major organs (H, heart; L, liver; S, spleen; Lu, lung; K, kidney) and tumor (T) harvested 24 h post-intravenous administration. (C) Heatmap visualization of fluorescence signal distribution within tumor regions.
(D) Semi-quantitative analysis of organ-specific fluorescence intensity. (E) Hemolysis analysis of blood samples incubated with CuTF-DC_AC50 at varying concentrations, using PBS and water as negative and positive controls, respectively. Inset: Hemolysis photographs of CuTF-DC_AC50 with different concentrations.
(F) Hematological and (G) biochemical analysis performed at 14 d post-injection. (H) Histopathological micrographs of H&E-stained mouse organ slices harvested upon treatment termination.Fig. 8In vivo antitumor therapeutic efficacy.
(A) Diagram depicting in vivo therapeutic potency assessment in CT26 tumor models. (B) Corresponding tumor photographs on day 14 post-treatment. Scale bar: 1 cm.
(C) Dynamic volumetric growth profiles of tumors among treatment cohorts. (D) Ex vivo tumor weight changes in distinct cohorts. (E) Individual tumor growth profiles among mice of each cohort (n = 5).
(F) Body weight changes of mice monitored throughout the therapeutic duration. (G) Immunohistochemical detection of post-treatment tumor tissues stained with H&E, Ki67, and TUNEL. (H) Immunofluorescence staining of tumor tissues for DLAT, c-caspase-1, GSDMD-NT, CD4&CD8 following the treatments.
Data are shown as mean ± SD (n = 3). p values were derived from a one-way ANOVA test, **p < 0.01, ***p < 0.001.
A critical step preceding the assessment of in vivo therapeutic efficacy was the evaluation of the biosafety profile for CuTF-DC_AC50. Hemocompatibility assessed via hemolysis assay revealed negligible hemolysis (rates < 5%) over the concentration ranging from 0 to 200 μg mL−1, confirming excellent blood compatibility (Fig. 7E).
Furthermore, after 14 d of consecutive administration, hematological and biochemical analyses of blood samples showed no significant alterations in key parameters relative to the blank cohort (Fig. 7F–G). H&E staining was conducted for histopathological observation of key visceral organs, including the heart, liver, spleen, lung, and kidney.
No obvious inflammatory infiltration or histological impairment was observed in these tissue samples (Fig. 7H). Taken together, these results underscore the excellent biocompatibility and favorable biosafety profile of CuTF-DC_AC50, strongly supporting its potential for further therapeutic translation.
Accordingly, to assess the in vivo tumor-suppressive potency, BALB/c mice bearing CT26 neoplasms were divided into five experimental cohorts: G1. Control; G2. DC_AC50; G3.
CuTF; G4. CuTF-DC_AC50; and G5. CuTF-DC_AC50 + αPD-L1.
Over the 14 consecutive days of treatment (Fig. 8A), no evident declines in body weight were recorded, corroborating the satisfactory systemic biocompatibility (Fig. 8F).
Tumor proliferation was markedly inhibited in the CuTF-DC_AC50 cohort, while the strongest antitumor effect was observed in the combination group (CuTF-DC_AC50 + αPD-L1) (Fig. 8B, C and E). Consistent with these results, the combination therapy showed a pronounced reduction in final tumor mass, representing only 23.5% of the average weight measured in the control group (Fig.
8D). Histological analyses further corroborated the therapeutic efficacy: H&E and TUNEL assays demonstrated severe tumor cell injury in the combination group, while Ki67 staining displayed a pronounced suppression of proliferation (Fig. 8G).
To elucidate the underlying mechanisms in vivo, immunofluorescence assays were carried out on tumor sections. The findings validated that CuTF-DC_AC50 administration could effectively trigger DLAT aggregation and elevate the levels of c-caspase-1 and GSDMD-NT (Fig. 8H).
In summary, CuTF-DC_AC50 exhibits excellent biosafety and efficient antitumor efficacy by simultaneously triggering cuproptosis and pyroptosis to suppress tumor proliferation in vivo. Notably, the combination with αPD-L1 ICB therapy further amplified the antitumor efficacy, highlighting a promising combinatorial strategy for cancer immunotherapy.
Transcriptomic analysis reveals therapeutic mechanism of CuTF-DC_AC50
To comprehensively elucidate the molecular-level antitumor mechanism of CuTF-DC_AC50, we employed transcriptome sequencing (RNA-seq) on CT26 cells following treatment. Systematic comparison of gene expression profiles across control and CuTF-DC_AC50-administered cohorts revealed high intra-group consistency and a clear separation in principal component analysis (PCA), indicating profound transcriptome remodeling induced by CuTF-DC_AC50 (Fig. 9A–B).
Venn analysis of the transcriptomic profiles screened out a transcriptome of 10719 genes that were uniformly expressed across both groups. In contrast, 1111 and 1548 transcripts were exclusively detected in the control and CuTF-DC_AC50-treated groups, respectively (Fig. 9C).
Differential expression analysis obtained 8686 differentially expressed genes (DEGs), with 5405 upregulated and 3281 downregulated (Fig. 9D–F), underscoring the extensive impact of CuTF-DC_AC50 on the cellular transcriptome.Fig. 9RNA sequencing analysis of the therapeutic mechanisms of CuTF-DC_AC50 in CT26 cells.
(A) Heatmap of the inter-sample correlation analysis across the control and CuTF-DC_AC50 cohorts. (B) PCA of transcriptomes from the control and CuTF-DC_AC50 cohorts. (C) Venn diagram depicting shared and unique expressed genes across the control and CuTF-DC_AC50 cohorts.
(D) Bar graph displaying counts of upregulated and downregulated DEGs in the two groups. (E) Volcano plot visualizing the distribution of DEGs. (F) Clustering heatmap visualizing the DEGs of the control and CuTF-DC_AC50 treatment cohorts.
(G) KEGG pathway enrichment statistics for DEGs derived from the two cohorts. (H) Multidimensional circular visualization of GO enrichment for the DEGs identified within the CuTF-DC_AC50 cohort relative to the control. (I) Chord diagrams of GO enrichment analysis.
(J-M) GSEA profiling functional variations between the experimental groups. (N-P) Heatmaps depicting the expression patterns of cuproptosis-, pyroptosis-, and immunity-related DEGs across different cohorts. (Q) Protein−protein interaction network analysis of genes linked to cuproptosis.
Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis highlighted remarkable alterations in key processes including endoplasmic reticulum protein processing, the NOD-like receptor signaling pathway, the TNF signaling pathway, oxidative phosphorylation, and the FoxO signaling pathway (Fig. 9G and Fig. S35).
Gene Ontology (GO) enrichment further confirmed these observations, showing predominant enrichment of DEGs in biological processes (BP), cellular components (CC) and molecular functions (MF), with BP representing the highest proportion (Fig. 9H). Chord diagrams and Cleveland dot plots visualized gene expressions associated with cellular BP under CuTF-DC_AC50 treatment, such as mitochondrial respiratory chain complex assembly, regulation of metal ion transport and immune effector processes, suggesting a coordinated multi-pathway response (Fig.
9I and Fig. S36). Gene Set Enrichment Analysis (GSEA) provided deeper insights into specific pathway modulation.
Upon treatment with CuTF-DC_AC50, rapid intracellular copper accumulation disrupts metal ion homeostasis, as evidenced by the significant activation of the "regulation of metal ion transport" pathway (Fig. 9J). Excess copper induces mitochondrial dysfunction by impairing "oxidative phosphorylation" (Fig.
9K), thereby triggering oxidative stress and initiating cuproptosis. Concurrently, suppression of the "negative regulation of NF-κB transcription factor activity" indicates enhanced NF-κB signaling (Fig. 9L), which promotes a pro-inflammatory microenvironment.
This environment facilitates pyroptosis and the release of DAMPs, thereby activating the "TNF signaling pathway" and stimulating ICD (Fig. 9M). Collectively, this cascade culminates in immune activation within the TME, linking copper-induced cytotoxic stress to downstream inflammatory and immune responses.
Clustering heatmaps of genes related to cuproptosis, pyroptosis, and immune response confirmed significant expression alterations in the CuTF-DC_AC50-treated group (Fig. 9N–P). Specifically, CuTF-DC_AC50 mediated an effective induction of cuproptosis, as reflected by a substantial reduction in the transcript levels of pivotal genes governing this cell death process.
Meanwhile, the treatment significantly upregulated pyroptosis- and immune-related genes. Among these, IL-6, a pivotal pro-inflammatory cytokine, serves as a core biomarker for amplified inflammatory responses triggered by pyroptosis [57], [58], [59]. JAK1, a key component of the JAK-STAT signaling pathway, functions as an upstream regulator of PD-L1 expression [60], [61], [62].
The coordinated changes in these genes confirm that CuTF-DC_AC50 effectively triggers cuproptosis and pyroptosis, and elicits a robust antitumor immune response. The analysis of a protein-protein interaction (PPI) network established from the DEGs disclosed significant functional connectivity, emphasizing their synergistic influence on key cellular mechanisms (Fig. 9Q and Fig.
S37). Taken together, these transcriptomic findings illustrate that CuTF-DC_AC50 exerts its potent antitumor effect by orchestrating a multifaceted response involving induction of cuproptosis, activation of pyroptosis, and elicitation of ICD.
Conclusion
In conclusion, we engineered CuTF-DC_AC50, an innovative copper homeostasis nanomodulator leveraging a unique "threefold dual" strategy for advanced cancer immunotherapy. Upon dual activation by the acidic and GSH-rich TME, this nanomodulator initiates a coordinated therapeutic cascade. It induces severe intracellular copper overload through a "dual-channel" approach: simultaneously releasing Cu2+ and inhibiting ATOX1-mediated Cu+ efflux, while disrupting the cellular antioxidant barrier via dual GSH depletion mechanism.
Consequently, this targeted copper surge triggers two cell death modalities, directly inducing cuproptosis via mitochondrial dysfunction and instigating pyroptosis via the ROS-activated caspase-1/GSDMD axis.
The synergistic induction of cuproptosis and pyroptosis potently drives ICD. This process effectively remodels the immunosuppressive TME from "cold" toward an immune-active "hot" state, characterized by promoted DC maturation, increased CD8+ T cell infiltration, and M2-to-M1 macrophage repolarization. Moreover, based on the finding that the nanomodulator upregulates tumor PD-L1 expression, we evaluated its combination with αPD-L1 ICB in vivo.
The outcomes confirmed that combined CuTF-DC_AC50 and αPD-L1 therapy yielded profound synergistic antitumor efficacy, achieving a remarkable 78.7% tumor growth inhibition that significantly outperformed monotherapies. Overall, by elegantly orchestrating precise TME modulation, dual-pathway cell death, and ICB therapy, this work establishes a robust and highly translatable combinatorial framework for next-generation cancer immunotherapy.
Materials and methods
Materials
CuCl2·2H2O were purchased from Macklin (Shanghai, China). DC_AC50 was purchased from Shanghai Yuanye Bio-Technology Co., Ltd. (Shanghai, China).
Pluronic F127, Glutathione (GSH), 5,5′-dithiomethyl (2-nitrobenzoic acid) (DTNB), and Methylene Blue (MB) were purchased from Aladdin (Shanghai, China). CCK8 detection kit, Reactive Oxygen Species (ROS) assay kit, Calcein AM/propidium iodide (PI) live/dead cell double staining kit were purchased from Solarbio Science & Technology Co., Ltd. (Beijing, China).
GSH content assay kit, mitochondrial membrane potential detection kit (JC-1), Annexin V-FITC/PI apoptosis detection kit and MitoTracker Red CMXRos were purchased from Servicebio Technology Co., Ltd. (Wuhan, China). Lactate dehydrogenase cytotoxicity detection kit, ATP content detection kit and Lyso-Tracker Red were purchased from Beyotime Biotechnology (Shanghai, China).
FITC-anti-CD11b, FITC-anti-CD11c, APC-anti-CD80, PE-anti-CD86, APC-anti-CD206, PerCP/Cy5.5-F4/80, PerCP-anti-CD45, FITC-anti-CD3, APC-anti-CD4, PE-anti-CD8, and PE-anti-FOXP3 were purchased from BioLegend, Inc. (San Diego, USA). IL-1β, TNF-α, IFN-γ, IL-10, IL-6 assay kit (ELISA) were purchased from Dogesce (Beijing, China).
BCA protein assay kit, high-sensitivity enhanced chemiluminescence (ECL) detection kit, and protein marker were purchased from SEVEN Biotech Co., Ltd. (Beijing, China). Primary antibodies for DLAT and FDX1 used in Western blot analysis were purchased from Abcam Co., Ltd.
(Shanghai, China). ATOX1 antibody was purchased from Proteintech Co., Ltd. (Wuhan, China).
PD-L1 antibody was purchased from Abmart Co., Ltd. (Shanghai, China), Caspase-1 and GSDMD antibodies were purchased from Cohesion Biosciences Co., Ltd. (Suzhou, China).
Characterizations
Transmission electron microscopy (TEM) and high-resolution TEM (HR-TEM) images were acquired using a JEOL JEM-2100F field-emission transmission electron microscope. Particle size distribution and zeta potential were measured by dynamic light scattering (DLS) with a Malvern Zetasizer Nano ZS90. Nitrogen adsorption-desorption isotherms were measured using an Autosorb-iQ gas sorption analyzer (Quantachrome Instruments, USA).
X-ray photoelectron spectroscopy (XPS) analysis was conducted on a Thermo VG Scientific ESCALAB250 spectrometer. Fourier transform infrared (FT-IR) spectra were collected on a Bruker Vertex Perkin-Elmer 580BIR spectrophotometer with KBr pellets. Steady-state and time-resolved fluorescence spectra were recorded on an FLS1000 spectrofluorometer (Edinburgh Instruments, UK).
UV-visible absorption spectra were recorded on a Hitachi U-3900 spectrophotometer. Elemental composition was determined using a PerkinElmer NexION 5000 inductively coupled plasma mass spectrometer (ICP-MS). Biological transmission electron microscope images were acquired with a Hitachi HT7700 transmission electron microscope.
Immunofluorescence images were acquired with an FV3000 laser scanning confocal microscope (Olympus Corporation, Japan). Flow cytometry analysis was performed on a Navios flow cytometer (Beckman Coulter, Ireland) equipped with 3 lasers and capable of 10-color detection. In vivo fluorescence imaging of mice was performed using an NIR-II fluorescence imaging system (Suzhou NIR-Optics Co., Ltd., Suzhou, China).
Preparation of CuT and CuT-DC_AC50 NPs
0.2 mL of 0.4 M CuCl2 aqueous solution and 0.6 mL of DMSO with or without 0.2 mg DC_AC50 were sequentially added to 4 mL of 0.04 M TPH aqueous solution under vigorous stirring. Subsequently, the mixed solution was maintained at 37 °C for 12 h with shaking at 100 rpm. The resulting product was washed three times with deionized water via centrifugation (12000 rpm, 10 min), then redispersed in ethanol to yield CuT and CuT-DC_AC50 NPs for further use.
Preparation of CuTF-DC_AC50 NPs
CuT-DC_AC50 dispersed in ethanol was mixed with F127 at a ratio of 1:3. After 10 min of sonication in ice bath, the mixed solution was magnetically stirred at ambient temperature for 2 h. Following ethanol removal via rotary evaporation, CuTF-DC_AC50 was dispersed in deionized water for further use.
Extracellular GSH depletion
The concentration-dependent GSH depletion capability of CuTF-DC_AC50 was evaluated. Briefly, 1.5 mL of CuTF-DC_AC50 with gradient concentrations (0, 25, 50, 100, 200, 300 μg/mL) were co-incubated with 10 μL 100 mM GSH at 37 °C for 60 min. Subsequently, 100 μL of each assay solution was collected and mixed with 1.5 mL of DTNB (50 μg/mL).
UV-Vis spectroscopy was applied to determine the optical density of TNB at 412 nm.
The time-dependent GSH depletion was further investigated. A mixture containing 1.5 mL of CuTF-DC_AC50 (100 μg/mL) and 10 μL of 100 mM GSH was incubated at 37 °C. Samples of 100 μL were harvested at designated time points and mixed with 1.5 mL DTNB (50 μg/mL).
UV-Vis spectroscopy was applied to determine the optical density of TNB at 412 nm.
Extracellular •OH detection
To evaluate the time dependence of •OH generation, a reaction mixture was prepared containing CuTF-DC_AC50 (300 μg/mL), GSH (2 mM), H2O2 (1 mM), and methylene blue (MB, 5 μg/mL) in a weakly acidic buffer (pH 6.5). With continuous stirring at 37 °C, MB degradation was tracked through measuring the reduction in absorbance at 664 nm using UV-Vis spectrometry at preset time points.
To evaluate the concentration dependence of •OH generation, CuTF-DC_AC50 at gradient concentrations (0, 50, 100, 200, 300 μg/mL) was incubated in weakly acidic buffer (pH 6.5). Each reaction mixture contained fixed concentrations of GSH (2 mM), H2O2 (1 mM), and MB (5 μg/mL). After a 2 h incubation at 37 °C with stirring, •OH generation was assessed by monitoring the decrease in the optical absorption of MB at 664 nm.
Degradation and release of CuTF-DC_AC50
To investigate the stimuli-responsive degradation characteristics of CuTF-DC_AC50, the nanomodulator were incubated under different conditions (i.e., PBS at pH 7.4, pH 6.5, pH 7.4 with GSH, and pH 6.5 with GSH) at 37 °C under shaking at 100 rpm for 24 h. Subsequently, TEM was employed to observe morphological variations of the nanomodulator.
To determine the release profile quantitatively, 10 mg of CuTF-DC_AC50 were suspended in 10 mL of PBS buffer under the aforementioned four conditions and incubated at 37 °C under shaking at 100 rpm. Upon reaching the set time points, 2 mL aliquots were withdrawn and centrifuged. The supernatant was harvested for the quantification of Cu2+ by ICP-MS and released DC_AC50 by fluorescence spectrometry.
A constant volume was maintained throughout the experiment by adding fresh buffer equal to the sample volume after each collection.
Cell culture
CT26 cells were grown in complete RPMI 1640 medium containing 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin at 37 °C under 5% CO2 humidified conditions. NIH3T3 cells were grown in complete DMEM medium containing 10% FBS and 1% penicillin-streptomycin at 37 °C under 5% CO2 humidified conditions.
Cellular uptake
For CLSM observation of cellular uptake profiles, CT26 cells were plated on culture slides and incubated for 24 h. Afterwards, the cells were treated with FITC-labeled CuTF-DC_AC50 at 200 μg/mL for different incubation periods. Subsequent to incubation, the cells underwent three PBS washes and were immobilized with 5% paraformaldehyde for 15 min.
After fixation, cell nuclei were stained with DAPI over 3 min, prior to three further PBS washes. The slides were mounted using antifade mounting medium and imaged by CLSM.
Lysosomal co-localization
After being seeded onto culture slides, CT26 cells were allowed to adhere for 24 h. Following this, the cells were cultured with FITC-labeled CuTF-DC_AC50 (200 μg/mL) at designated time points (4, 6, and 8 h). Post-treatment, the cells were rinsed thrice with PBS, then stained with LysoTracker™ Red and incubated at 37 °C in darkness for 60 min.
Following staining, three additional PBS washes were performed. The slides were mounted using antifade reagent and visualized by confocal laser scanning microscope (CLSM).
Cytotoxicity measurement
To verify the cellular safety of CuTF-DC_AC50, CT26 and NIH3T3 cells were plated into 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Different concentrations of CuTF-DC_AC50 were then applied for another 24 h. After adding CCK-8 reagent, the optical density at 450 nm was determined using a microplate reader.
To assess cellular viability under different conditions, CT26 cells were plated into 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Different concentrations of CuTF-DC_AC50 were then applied for another 24 h in distinct conditions (pH 7.4 or pH 6.5 and with or without 5 mM GSH). After adding CCK-8 reagent, the optical density at 450 nm was determined using a microplate reader.
To assess cellular viability under different formulations, CT26 cells were plated into 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. The cells were further incubated with different concentrations of the formulations (CuTF and CuTF-DC_AC50: 0-200 μg/mL; corresponding DC_AC50 concentrations: 0-10 μg/mL) for another 24 h in pH 6.5 medium containing 5 mM GSH. After adding CCK-8 reagent, the optical density at 450 nm was determined using a microplate reader.
Live/dead cell staining
Cell viability was assessed using calcein-AM/PI staining. CT26 cells were plated into 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Subsequently, the cells were incubated with different formulations (control: PBS; DC_AC50: 10 μg/mL; CuTF and CuTF-DC_AC50: 200 μg/mL) for another 24 h.
After triple PBS washing, the cells were subjected to calcein-AM/PI staining for 30 min, then observed and photographed under an inverted fluorescence microscope.
Colony formation assay
CT26 cells were plated into 6-well plates at 1 × 103 cells/well and incubated for 72 h. Thereafter, the cells were exposed to various formulations over 24 h. The culture medium was refreshed, and the cells were further cultivated for 7 days before staining with 0.2% crystal violet.
Apoptosis analysis
Annexin V-FITC/PI dual staining combined with flow cytometry was used to detect cell apoptosis. CT26 cells were plated in 6-well plates at 3 × 105 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were digested with EDTA-free trypsin and centrifuged at 300×g for 5 min.
After rinsing with PBS, cells sediments were resuspended in binding buffer. After 15 min of dark incubation with the two fluorescent dyes, all samples were analyzed via flow cytometry.
Intracellular GSH assay
CT26 cells were plated in 6-well plates at 3 × 105 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were rinsed with PBS and harvested via centrifugation. Intracellular GSH content was then determined with a GSH detection kit.
Intracellular ROS detection
For ROS detection, CT26 cells were plated in 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were rinsed with PBS. Subsequently, the cells were stained with DCFH-DA probe and kept at 37 °C in darkness for 30 min, prior to additional PBS rinses.
Relevant fluorescent signals were visualized and photographed via an inverted fluorescent microscope.
For flow cytometry detection, CT26 cells were plated in 6-well plates at 3 × 105 cells/well and pre-cultured for 24 h. After treatment for 24 h with designated experimental groups, the cells were digested with trypsin, collected via centrifugation and transferred to microcentrifuge tubes. Following 30 min dark incubation with DCFH-DA at 37 °C, the cells were rinsed and resuspended in PBS.
Intracellular ROS levels were then quantified via flow cytometry.
Mitochondrial membrane potential assay
CT26 cells were plated in 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were rinsed with PBS. After 30 min dark staining with JC-1 working solution at 37 °C, the cells were subjected to additional PBS washes, and fluorescent photographs were recorded via an inverted fluorescence microscope.
Mitochondrial staining
CT26 cells were plated on culture slides and incubated for 24 h. Upon 24 h of intervention with various formulations, the cells were stained with MitoTracker Red CMXRos and incubated at 37 °C for 15 min under dark conditions. After staining, three additional PBS washes were performed.
CLSM was used to capture images of the cell slides.
Immunofluorescence staining
CT26 cells were plated on culture slides and incubated for 24 h, followed by co-incubation with various formulations for another 24 h. After being fixed in 5% paraformaldehyde, the cells were permeabilized using 0.2% Triton X-100 and blocked with 5% serum. The cells were incubated with primary antibodies overnight, followed by 1 h incubation with Alexa Fluor 488-labeled secondary antibodies.
Cell nuclei were counterstained with DAPI. Afterwards, slides were sealed with anti-fade mounting medium and visualized via CLSM.
Western blot
CT26 cells were plated in 6-well plates at 3 × 105 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were lysed on ice, and centrifuged to collect supernatants. After protein quantification, samples were boiled for denaturation.
After loading denatured proteins into SDS-PAGE gels and completing electrophoretic transfer to PVDF membranes, samples were blocked with 5% skim milk for 2 h at room temperature. Membranes were exposed to primary antibodies overnight at 4 °C, followed by a 2 h incubation with the corresponding secondary antibodies at ambient temperature. Protein bands were visualized with chemiluminescent substrate reagent.
Lactate dehydrogenase (LDH) assay
CT26 cells were plated in 96-well plates at 1 × 104 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, LDH levels were measured via LDH cytotoxicity detection kit.
Measurement of intracellular ATP
CT26 cells were plated in 6-well plates at 3 × 105 cells/well and pre-cultured for 24 h. Upon 24 h of intervention with various formulations, the cells were harvested and intracellular ATP content was determined using ATP detection kits.
CuTF-DC_AC50-induced BMDC maturation in vitro
For in vitro evaluation of DC maturation, mouse bone marrow cells were incubated with GM-CSF (20 ng/mL) and IL-4 (10 ng/mL) over 10 days to obtain bone marrow-derived dendritic cells (BMDCs). A 6-well transwell system with a 0.4 μm pore size was employed for the BMDC maturation assay. CT26 cells from the various treatment groups were seeded in the upper chamber, while BMDCs were plated in the lower chamber.
After 24 h of co-culture, BMDCs were collected. The expression levels of CD80 and CD86 were detected by flow cytometry.
Animal models
Eight-week-old female BALB/c mice (18–20 g) were supplied by Weitong Lihua Experimental Animal Technology Co., Ltd (Beijing). All animal experiments were performed following the guidelines of the Experimental Animal Center of Shanxi Medical University and approved by the Institutional Animal Care and Use Committee (IACUC) of the university (Approval No. 2024-0007).
For tumor model construction, CT26 cells resuspended in 50 μL PBS were subcutaneously inoculated into the right posterior flank of mice. In vivo experiments were initiated once tumor volume grew to around 80 mm3.
Evaluation of antitumor immunity
CT26 tumor-bearing mice were randomly assigned into five groups (n = 5) for corresponding treatments: (1) control (PBS); (2) DC_AC50 (300 μg/kg); (3) CuTF (6 mg/kg); (4) CuTF-DC_AC50 (6 mg/kg); (5) CuTF-DC_AC50 + αPD-L1 (CuTF-DC_AC50: 6 mg/kg; αPD-L1: 8 mg/kg). PBS, DC_AC50, CuTF, CuTF-DC_AC50 were administered via tail vein injection on days 0, 3, 6, and 9, while αPD-L1 was delivered intraperitoneally on days 1, 4, 7, and 10. All mice were euthanized on day 14.
Fresh spleen, lymph nodes, and tumor tissues were collected and processed into single-cell suspensions. Fluorescence-tagged antibodies were used to stain the isolated cells, followed by flow cytometry detection.
Hemolysis analysis
Blood was collected from BALB/c mice into anticoagulant-containing tubes and gently mixed. The anticoagulated blood was then centrifuged at room temperature at a relative centrifugal force of 300-500 × g for 5-10 min. After centrifugation, the upper plasma supernatant was gently aspirated and discarded.
After washing the red blood cell (RBC) pellet with PBS for three cycles, 5 mL PBS was supplemented to resuspend erythrocytes. Then, different concentrations of CuTF-DC_AC50 were co-incubated with 300 μL of RBC suspension at 37 °C with shaking at 100 rpm for 3 h. PBS and deionized water were set as negative and positive controls, respectively.
Finally, the absorbance at 542 nm was determined using a microplate reader.
In vivo fluorescence imaging
To monitor the in vivo biodistribution of CuTF-DC_AC50, IR820-labeled CuTF-DC_AC50 (6 mg/kg) were administered into CT26 tumor-bearing mice through the tail vein. Fluorescence signals in mice were captured at various time points via a NIR-II imaging platform. In addition, main visceral organs including heart, liver, spleen, lung, kidney as well as tumor tissues were harvested at 24 h after CuTF-DC_AC50 injection for ex vivo fluorescent imaging.
Evaluation of antitumor efficacy
CT26 tumor-bearing mice were randomly assigned into five groups (n = 5) for corresponding treatments: (1) control (PBS); (2) DC_AC50 (300 μg/kg); (3) CuTF (6 mg/kg); (4) CuTF-DC_AC50 (6 mg/kg); (5) CuTF-DC_AC50 + αPD-L1 (CuTF-DC_AC50: 6 mg/kg; αPD-L1: 8 mg/kg). PBS, DC_AC50, CuTF, CuTF-DC_AC50 were administered via tail vein injection on days 0, 3, 6, and 9, whereas αPD-L1 was delivered intraperitoneally on days 1, 4, 7, and 10. Tumor growth and mouse body weight were monitored every two days.
Tumor volume was computed via the following formula: Volume=(Length)×(Width)2/2. All mice were euthanized on the 14th day after treatment. Isolated tumor tissues were weighed and subjected to histological examinations including H&E, TUNEL and Ki67 staining.
Main organs were harvested for H&E staining to assess systemic biosafety.
Statistical analysis
Data were presented as the mean ± standard deviation. One-way ANOVA test was used to evaluate the statistical significance. Differences were considered statistically significant at *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.
CRediT authorship contribution statement
Qian Wang: Writing – original draft, Methodology, Data curation, Conceptualization. Qian Wang: Methodology, Formal analysis, Data curation. Jie Dong: Validation, Data curation, Conceptualization.
Yuxin Zhou: Methodology, Investigation, Formal analysis. Zichen Cui: Visualization, Investigation. Baole Li: Visualization, Investigation.
Botao Qu: Writing – review & editing, Funding acquisition, Conceptualization. Ruiping Zhang: Writing – review & editing, Supervision, Funding acquisition.
Declaration of competing interest
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.