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. 2024 Sep 2;13(31):2402349. doi: 10.1002/adhm.202402349

An Oxidative Stress Nano‐Amplifier for Improved Tumor Elimination and Combined Immunotherapy

Wei Zhang 1, Yijun Ran 2, Mi Yang 1, Yaqin Hu 1, Zhigang Wang 1, Yang Cao 1, Haitao Ran 1,✉
PMCID: PMC11650535  PMID: 39221686

Abstract

Amplifying oxidative stress to disrupt intracellular redox homeostasis can accelerate tumor cell death. In this work, an oxidative stress amplifier (PP@T) is prepared for enhanced tumor oxidation therapy to reduce tumor growth and metastases. The nano‐amplifier has been successfully constructed by embedding MTH1 inhibitor (TH588) in the PDA‐coated porphyrin metal–organic framework PCN‐224. The controllable‐released TH588 is demonstrated from pores can hinder MTH1‐mediated damage‐repairing process by preventing the hydrolysis of 8‐oxo‐dG, thereby amplifying oxidative stress and exacerbating the oxidative DNA damage induced by the sonodynamic therapy of PP@T under ultrasound irradiation. Furthermore, PP@T can effectively induce immunogenic cell death to trigger systemic anti‐tumor immune response. When administered in combination with immune checkpoint blockade, PP@T not only impedes the progression of the primary tumor but also achieves obvious antimetastasis in breast cancer murine models, including orthotopic and artificial whole‐body metastasis models. Furthermore, the nanoplatform also provides photoacoustic imaging for in vivo treatment guidance. In conclusion, by amplifying oxidative stress and reactive oxygen species sensitized immunotherapy, this image‐guided nanosystem shows potential for highly specific, effective combined therapy against tumor cells with negligible side‐effects to normal cells which will provide a new insight for precise tumor treatment.

Keywords: immunogenic death, MutT homologue 1 (MTH1), oxidative stress, reactive oxygen species (ROS), sonodynamic therapy (SDT)

An oxidative stress amplifier (PP@T) is developed for more efficient tumor therapy by amplifying oxidative stress and reactive oxygen species sensitized immunotherapy, and have conducted extensive research on anti‐tumor mechanisms.

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1. Introduction

Cancer has a much higher fatality rate than other diseases worldwide, and it is one of the most destructive and profoundly disrupting illnesses.[ 1 ] It is a major issue across the world to develop an effective anticancer therapy. Oxidative stress (OS), characterized by an imbalance in redox favoring oxidant burden, is a general phenotype of many cancers and plays a crucial role in the development, progression, and spread of cancer.[ 2 ] Cancer cells typically exhibit higher levels of oxidants, which suggests a dual therapeutic strategies by regulating redox status (i.e., pro‐oxidant therapy and/or antioxidant therapy).[ 3 ] This knowledge and understanding of the altered redox status of cancer cells has inspired many researchers to develop new cancer treatment strategies. Among various signaling molecules influencing cancer cells, reactive oxygen species (ROS) have been emerged as key players underlying the acquisition of multiple hallmarks.[ 4 ] As the role of ROS in various stages of tumor development, such as early tumor occurrence, malignant progression, invasion, and metastasis, has been clarified, its application in tumor therapy has attracted more and more attention.[ 5 ] Neoplastic cells' ROS levels can rise directly through ROS generation or indirectly through disruption of the antioxidant cascade.[ 6 ] While it seems reasonable to explore ways to disrupt ROS homeostasis in cancer cells by decreasing or increasing ROS levels, the latter dominates, as the results obtained by the former, particularly antioxidants, are currently controversial.

The regulation of dysfunctional redox and an increased ROS tension can lead directly to oxidative damage of DNA, or to oxidative damage of free bases in the cellular and mitochondrial deoxyribonucleoside triphosphate (dNTP) pools.[ 7 ] However, DNA damage repair mechanisms in tumor cells can compromise the therapeutic efficacy of oxidative therapy in tumor treatment.[ 8 ] The MutT homolog 1 (MTH1) protein sanitizes the oxidized dNTP pool, by converting 2‐OH‐dATP or 8‐oxod‐GTP into 2‐OH‐dAMP or 8‐oxod‐GMP, thereby avoiding incorporation of these oxidized nucleotides into the DNA, which could otherwise lead to mismatch, mutations and cell death.[ 9 ] Given that the MTH1 protein is thought to minimize cancer‐related damage and sterilize the oxidized dNTP pool, it has been reported that inhibiting the DNA damage repair pathway regulated by MTH1 may lead to increased oxidative damage and cell death, thereby enhancing therapeutic efficacy.[ 10 ] To take advantage of MTH1's effectiveness as an anticancer target for oxidative treatment, a family of MTH1 inhibitors has recently been developed.[ 7 , 11 ] Once coupled to the active site of MTH1, the clearance of 8‐oxo‐dG can be effectively hindered by this inhibitor and induce apoptosis in many types of cancer cells.[ 12 ] However, such oxidative therapy relies on endogenous ROS to produce 8‐oxo‐dG.[ 13 ]

Sonodynamic therapy (SDT) is a recently developed new approach to cancer treatment that relies on ultrasound (US) combined with a substance that responds to US to produce ROS.[ 14 ] Due to its remarkable tissue penetration depth and minimal side effects of external ultrasound waves, SDT has gained significant attention as a promising strategy for cancer treatment.[ 15 ] Although the main mechanism of SDT‐induced apoptosis is the production of reactive oxygen species rather than the direct damage of DNA, the dNTP pool oxidation may be caused during SDT. Since free dNTP is more sensitive to ROS than DNA double‐stranded,[ 16 ] it can be envisioned that SDT could act as a source of ROS along with MTH1 inhibitors, producing more 8‐oxo‐dG to kill cancer cells.

Herein, an amplified oxidative damage strategy is proposed for tumor therapy, which focuses not only on ROS production, but also on inhibiting subsequent MTH1 enzyme activity in tumor cells (Figure  1 ). The nano‐amplifier (PP@T), were utilized porphyrin‐based PCN‐224 as the sonosensitizer, which also serves as a nanocarrier for delivering the MTH1 inhibitor (TH588). Polydopamine (PDA) was coated as the gatekeeper for stimulus‐responsive drug release and photoacoustic (PA) imaging. The PP@T exhibited favorable dispersion stability and uniform nano‐size distribution. Under US irradiation, the PP@T generated singlet oxygen for SDT. Simultaneously, TH588 inhibited the activity of the MTH1 protein, preventing the hydrolysis of 8‐oxo‐dGTP and suppressing DNA damage repair.[ 7 , 17 ] Ultimately, SDT could work together with the MTH1 inhibitor as a source of ROS to produce more 8‐oxo‐dGTP in killing cancer cells, the amplification of oxidative stress resulting in superior therapeutic effects both in vitro and in vivo. Because both SDT and oxidative therapy could trigger immunogenic death, PP@T can also trigger more immunogenic death, which triggers an anti‐tumor immune response. PP@T can therefore be used as an adjuvant to carry out comprehensive anti‐tumor immunotherapy more effectively by inducing cell immunogenic death, promoting antigen presentation, and stimulating the immune system. When used in combination with immune checkpoint blocking, PP@T effectively inhibits the progression of the primary tumor and boost antitumor immune responses, thereby suppressing the development of tumor metastasis in murine models of 4T1 breast cancer. These models include both orthotopic and artificial whole‐body metastasis tumor models. Overall, this study suggests an overwhelming advantages of using oxidative stress amplification strategy to improve cancer immunotherapy, which will provide implications for the clinical development of combination therapies to block tumor growth and metastasis.

Figure 1.

Figure 1

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A) Preparation process of PP@T and B) Mechanism of PP@T for Tumor Cell Apoptosis and Amplified Oxidative Stress Activated Immunotherapy.

2. Results and Discussion

2.1. Synthesis and Characterization of PP@T

In this study, PCN‐224 was selected as the nanocarrier due to its high porphyrin loading capacity without experiencing self‐quenching, tunable surface properties, and nano‐porous channels.[ 18 ] The size of PCN‐224 could be finely adjusted by varying the amount of benzoic acid. Thus, a porous metallic organic framework, PCN‐224, was synthesized through a one‐step solvothermal method.[ 18 , 19 ] As depicted in Figure  2a, PCN‐224 nanoparticles were uniformly spherical, with an average size of ≈90 nm. Moreover, PCN‐224 nanoparticles possessed a BET surface area of 394.25 m2 g−1 and an average pore size of 4.54 nm, making them suitable for loading small molecule drugs (Figure 2c; Figure S1, Supporting Information). Subsequently, the MTH1 inhibitor TH588 was loaded into the PCN‐224 nanoparticles, with the drug loading ratio determined via high‐performance liquid chromatography (HPLC) to be 4.78%. The nanoparticles were further modified with polydopamine (PDA) to enhance biodegradability and ensure negligible long‐term toxicity. The surface modification with polydopamine changed the color of the nanoparticle solution from dark purple to black, as shown in Figure S2 (Supporting Information). Scanning electron microscopy (SEM) and TEM results demonstrated that PP@T nanoparticles maintained a spherical, uniform size with a smooth surface, resembling the original MOF templates. High‐angle annular dark‐field scanning transmission electron microscopy (HAADF‐STEM) was utilized to determine the chemical composition of PP@T and revealed that it mainly consisted of N, O, C, and Zr, with uniform distribution (Figure 2b; Figure S3a, Supporting Information). Dynamic light scattering (DLS) measurements (Figure 2d) indicated changes in particle size and surface potential distribution (Figure 2e) among different nanoparticles, confirming the successful PDA modification. UV‐vis spectra showed that PP@T, PP, and TCPP exhibited the same characteristic absorption peak at 413 nm (Figure 2f), confirming the synthesis of PP@T from PCN‐224. The released amount of TH588 in PCN‐224 after 48 h was ≈50%, which is about four times the release amount in PP@T (Figure S3b, Supporting Information). The differentiated release rates can be ascribed to the PDA coating obstructs the release of TH588 under physiological conditions.

Figure 2.

Figure 2

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Characterizations of the PP@T. a) TEM and SEM images of PCN‐224 and PP@T. The scale bar indicated 100 nm. b) Elemental mapping of PP@T. c) N2 adsorption‐desorption isotherms of PCN‐224. d,e) Size distribution and Zeta potentials of PCN‐224, PP, and PP@T. f) UV–vis spectrum of TCPP, PCN‐224, and PP@T. g) DPBF consumption of PP@T under LIFU irradiation. h) ESR spectra of 1O2 trapped by TEMP in PP@T dispersions upon LIFU irradiation for prolonged durations. Data were presented as the mean ± SD (n = 3).

Sonodynamic therapy assisted by nano‐sonosensitizers produces ROS, such as singlet oxygen, which can kill tumor cells.[ 20 ] To demonstrate the SDT capabilities of PP@T, the generation of ROS was quantitatively determined using 1,3‐diphenylisobenzofuran (DPBF).[ 21 ] The ethanol solution containing DPBF and PP@T exhibited a time‐dependent decrease in absorption intensity upon exposure to US irradiation, indicating the production of 1O2 by PP@T (Figure 2g). In contrast, when DPBF solution was treated with US alone, the intensity remained largely unchanged within 10 min, indicating that US alone had minimal impact on DPBF oxidation. Additionally, electron spin resonance (ESR) was employed to monitor the ROS species during the SDT process and to elucidate the mechanism of PP@T‐mediated SDT. As shown in Figure 2h, a strong and characteristic 1:1:1 signal was observed in the PP@T plus US group, with an even stronger signal observed with prolonged irradiation time, suggesting the production of 1O2. These results collectively suggest that PP@T are effective sonosensitizers capable of generating ROS following US irradiation.

The ability of PP@T to induce ROS production in tumor cells was assessed using 2′,7′‐dichlorofluorescein diacetate (DCFH‐DA) as a probe. DCFH‐DA does not fluoresce independently but is rapidly oxidized by ROS to form fluorescein 2′,7′‐dichlorofluorescein (DCF) within living cells, emitting green fluorescence.[ 22 ] As shown in Figure S4 (Supporting Information), 4T1 cells in the PP@T plus US group exhibited strong green fluorescence, whereas neither the pure ultrasound group nor the pure PP@T group displayed a significant fluorescence signal. Additionally, flow cytometry confirmed that PP@T effectively activated ROS production following ultrasound irradiation (Figure S5, Supporting Information), consistent with the results of confocal laser scanning microscopy.

2.2. Cellular Uptake of PP@T

To investigate the cellular uptake of PP@T by tumor cells, 4T1 cells were co‐incubated with the nanoparticles (50 µg mL−1). As shown in Figure S6 (Supporting Information), 4T1 cells exhibited time‐dependent uptake of PP@T after 1, 2, and 4 h of incubation. Furthermore, PP@T displayed a distinct cytoplasmic staining pattern of red fluorescence signal after incubation with 4T1 cells, indicating a gradual increase in nanoparticle uptake by tumor cells. Similarly, flow cytometry analysis was conducted to evaluate and quantify nanoparticle uptake by breast cancer cells (Figure S7, Supporting Information). The results revealed that the uptake rate of nanoparticles after 4 h of co‐incubation reached 81.58%, significantly higher than at other time points (23.35% and 58.32%) (p<0.05). The intracellular localization of PP@T was further examined through TEM, which revealed the presence of numerous nanoparticles within 4T1 cells (Figure S8, Supporting Information). No apparent morphological changes were observed after the cellular uptake of nanoparticles.

2.3. In Vitro Antitumor Effect of Oxidative Damage‐Enhanced SDT

To assess the antitumor effect of oxidative damage‐enhanced SDT, the cytotoxicity of PP@T was evaluated using a CCK‐8 assay. Initially, the effect of PP@T on the viability of HUVECs and 4T1 cells was assessed by the CCK‐8 assay. As depicted in Figure  3a, after 24 h of co‐incubation, no significant cytotoxicity was observed in 4T1 cells and HUVECs at various concentrations of PP@T. Even at a 200 µg mL−1 concentration, the cell viability of 4T1 cells and HUVECs remained above 85%, indicating the biocompatibility of PP@T.

Figure 3.

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a) Cell viability of 4T1 cells and HUVEC cells incubated with PP@T for 24 h. b) Cell viability of different samples against 4T1 cells with or without LIFU irradiation. (1: Control group, 2: LIFU group, 3: PP@T group, 4: PP+LIFU group, 5: PP@T+LIFU group) c) Plate clone formation assay of 4T1 cells treated with different conditions. d) Live/dead cell staining assay with different treatments against 4T1 cells. Dead cells were stained with PI (red), and live cells were stained with Calcein‐AM (green). Scale bar 50 µm. Data were presented as the mean ± s.d (n = 5). The level of statistical significance was indicated when appropriate (* P<0.05, ** P<0.01).

To evaluate the antitumor effect of SDT enhanced by oxidative damage, the cytotoxicity of PP@T was examined using a CCK‐8 assay. In the absence of TH588 loading, the viability of PP‐treated cells decreased with increasing concentration or power under low‐intensity focused ultrasound (LIFU) irradiation (Figure S9, Supporting Information), demonstrating the cytotoxic production of ROS by the nanoparticles. As shown in Figure 3b, nanoparticles loaded with TH588 (PP@T+LIFU group) exhibited significant cytotoxicity under ultrasound irradiation. In contrast, the PBS group, single LIFU group, and PP@T without ultrasound group displayed minimal cytotoxicity to 4T1 cells. Similarly, a partial cytotoxic effect was observed in the single SDT group (PP+LIFU). With the addition of TH588, the SDT efficacy of PCN‐224 was significantly enhanced due to the combined effect of ROS‐induced damage and inhibition of MTH1‐mediated DNA damage repair. Colony formation assays were also performed after different treatments, demonstrating that PP@T combined with LIFU significantly reduced the clonogenicity of 4T1 cells (Figure 3c), consistent with the results of the CCK‐8 assay. Confocal laser scanning microscopy images from live/dead cell staining further supported these findings. As shown in Figure 3d, the PP@T+ LIFU group exhibited the most effective tumor cell‐killing ability compared to the other treatment groups. Collectively, these results indicate that the MTH1 inhibitor TH588 sensitizes tumor cells to SDT, which can effectively amplify oxidative stress in tumor cells, resulting in a synergistic therapeutic effect.

2.4. The Photoacoustic (PA) Imaging and Biodistribution

The PA imaging capacity of PP@T was evaluated in vitro, confirming their effectiveness as contrast agents for PA imaging (Figures S10 and S11, Supporting Information). Subsequently, PA imaging was used to characterize the accumulation of PP@T at the tumor site. PA images of tumors were collected at different time points following intravenous injection of PP@T into 4T1 tumor‐bearing mice. As depicted in Figures S12 and S13 (Supporting Information), the PA signal from the tumor reached its peak intensity at 24 h post‐injection, indicating the accumulation of PP@T in tumors over time.

The biodistribution of nanoparticles was evaluated by measuring the fluorescence signal of PP@T labeled with 1,1′‐dioctadecyl‐3,3,3′,3′‐tetramethylindotricarbocyanine iodide (DIR) in various organs at different time points. As shown in Figures S14 and S15 (Supporting Information), nanoparticles gradually accumulated in tumors over time, reaching a peak at 24 h post‐injection, consistent with the PA imaging results. Moreover, the liver and spleen exhibited the highest nanoparticle absorption among major organs, suggesting that these organs primarily contributed to nanoparticle metabolism.

2.5. Mechanism Study of the Combined Therapy

To elucidate the mechanism underlying the reduction in cell viability induced by PP@T in 4T1 tumor cells, mitochondrial damage was assessed via flow cytometry using the JC‐1 mitochondrial membrane potential (MMP) assay kit. It is widely acknowledged that JC‐1 forms aggregates in the mitochondria of normal cells, whereas in apoptotic cells, JC‐1 accumulates as monomers in the cytoplasm due to MMP collapse.[ 23 ] As shown in Figure  4a, the PP@T+LIFU group exhibited a significantly higher JC‐1 monomer/aggregate ratio (53%) compared to the light control (3%) and PP@T alone (7%) groups. These results suggest that PP@T + LIFU disrupted the MMP in cells.

Figure 4.

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a) Mitochondrial membrane potential detection via JC‐1 staining. b) TEM images of 4T1 cells after different treatments. c) CLSM images of 8‐oxo‐dG, 4T1 cells were subjected with different treatments and co‐stain with DAPI and avidin‐Alexa Fluor 488. Scale bar 50 µm. Quantitative fluorescence analysis the colocalization of 8‐oxo‐dG with d) nuclei and e) entire cell by using Image (J). Data were presented as the mean ± s.d (n = 3). The level of statistical significance was indicated when appropriate (* P<0.05, ** P<0.01).

Subsequently, the rate of apoptosis induced by PP@T with US irradiation was investigated using the Annexin V‐FITC/PI staining assay. As demonstrated in Figure S16 (Supporting Information), the proportion of apoptotic cells was 6.17%, 7.15%, 21.20%, 45.97%, and 71.1% following treatment with PBS, LIFU, PP@T, PP + LIFU, and PP@T + LIFU, respectively. The PP@T + LIFU group exhibited the highest apoptosis ratio, consistent with the results of the CCK‐8 assay.

Furthermore, TEM was employed to evaluate the apoptotic effects induced by PP@T with US irradiation. As shown in Figure 4b, cells in the control group and the single ultrasound irradiation group exhibited normal size and shape, intact structures, and nuclei. However, cells treated with PP + LIFU displayed certain internal structural damage and destruction, particularly after treatment with PP@T + LIFU, where the internal structure of cells was completely disrupted. These cells displayed characteristic signs of apoptosis, such as membrane shrinkage, condensation of the cell nucleus, and the formation of phagosomes or apoptotic bodies.[ 24 ] These findings confirm that PP@T, in conjunction with US irradiation, induce tumor cell death through apoptosis.

In addition, the mechanism underlying the amplified oxidative DNA damage during combined therapy was explored. MTH1 is responsible for enzymatically converting oxidized nucleotides (8‐oxo‐dGTP) into their corresponding monophosphates, preventing cell damage and death.[ 25 ] Avidin is known to specifically bind to 8‐oxo‐dGTP. Cells subjected to different treatments were stained with Alexa Fluor 488‐conjugated Avidin and 4′,6‐diamidino‐2‐phenylindole (DAPI) to assess the incorporation of 8‐oxo‐dGTP into DNA. As depicted in Figure 4c–e, cells treated with PP@T + LIFU exhibited the highest accumulation of 8‐oxo‐dGTP, as evidenced by the enhanced green fluorescence of Avidin‐Alexa Fluor 488. Notably, the green signal of Avidin‐Alexa Fluor 488 overlapped with DAPI, indicating increased incorporation of 8‐oxo‐dGTP into nuclear DNA, inducing DNA mutations following PP@T‐SDT treatment. Moreover, the integrated intensity of green fluorescence in nuclear regions was quantified to assess DNA mutation. Cells treated with PP@T‐based SDT displayed 1.83 times higher 8‐oxo‐dGTP levels in nuclei than those treated with ultrasound alone. For the cells treated by the combined therapy (PP@T + LIFU group), displayed higher 8‐oxo‐dGTP levels than the PP@T treatment, revealing that the additional DNA mutant caused by SDT also capable of resulting in programmed cell death. These results demonstrate that PP@T can inhibit intracellular self‐repair processes by reducing the elimination of 8‐oxo‐dGTP, leading to the disruption of the ROS defense system in cancer cells and significantly enhancing oxidative damage induced by SDT.

2.6. Assessment of PP@T Augmented SDT for Suppression of Tumors

To evaluate the effectiveness of PP@T ‐augmented SDT, we employed a 4T1 xenograft Balb/c mice model. The experimental protocol is outlined in Figure  5a, with the mice divided into five groups: Control group, LIFU group, PP@T group, PP + LIFU group, and PP@T + LIFU group. After administering different treatments to each group, we assessed the therapeutic efficacy by monitoring changes in tumor volume. As depicted in Figure 5b–d, compared to the Control, LIFU, and NPs groups, mice treated with either PP combined with LIFU or PP@T combined with LIFU displayed effective tumor growth inhibition. Notably, individual SDT performed by PP exhibited partial tumor growth inhibition, while the PP@T combined with LIFU yielded significantly enhanced therapeutic outcomes. These findings were consistent with our in vitro results. The significant antitumor effect of PP@T can be attributed to the improved SDT efficiency of TH588. PP@T, once internalized, can inhibit the activity of MTH1, impeding the DNA damage repair process in cancer cells, thereby amplifying oxidative stress in tumor cells and enhancing the effectiveness of therapy.

Figure 5.

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a) Schematic illustration of the therapeutic immunization against orthotopic breast tumors. b) Tumor volume changes of mice after different treatments. c) Weights of tumors measured at the end of treatment. d) The photographs of tumors in different groups at the end of treatment. e) H&E, TUNEL, and PCNA staining of tumors (Scale bar 100 µm). Nuclei were stained with DAPI. f) Immunohistochemical analysis of 8‐oxo‐ dGTP and γH2AX (Scale bar 100 µm). Data were presented as the mean ± s.d (n = 5). The level of statistical significance was indicated when appropriate (* P<0.05, ** P<0.01).

In addition, we conducted H&E and immunohistochemistry staining on tumor tissues. The analyses, including H&E, TUNEL, and PCNA staining, revealed extensive cancer necrosis and apoptosis in the PP@T+LIFU treated group, indicating significant damage to the tumor tissue (Figure 5e). Immunofluorescence staining of MTH1 in tumor tissue confirmed reduced MTH1 expression after PP@T injection (Figure S17, Supporting Information). Moreover, the effects of TH588 treatment on 8‐oxo‐dGTP integration into DNA and DNA damage were evaluated through immunohistochemistry using 8‐oxo‐dGTP and γH2AX antibodies.[ 26 ] As illustrated in Figure 5f, TH588 significantly increased 8‐oxo‐dGTP integration and DNA damage in the PP@T + LIFU treated group compared to the other groups. Specifically, the PP@T + LIFU treated group exhibited a 3.46‐fold increase in 8‐oxo‐dGTP levels (Figure S18, Supporting Information) and a 1.69‐fold increase in γH2AX levels (Figure S19, Supporting Information) compared to the single SDT group. Finally, we assessed the in vivo biocompatibility of PP@T. Notably, PP@T did not induce changes in body weight (Figure S20, Supporting Information) or pathology (Figure S21, Supporting Information) in major organs in any experimental group, confirming its potential biocompatibility and safety.

2.7. Assessment of PP@T plus PD‐1 Blockade for Suppression of Distant Tumors

The above experimental results strongly support PP@T as an effective local therapeutic system for tumor treatment. However, cancer often leads to recurrence and metastasis.[ 27 ] Therefore, we explored the synergistic anti‐cancer therapeutic effect of PP@T‐enhanced SDT in combination with PD‐1 checkpoint blockade therapy for metastatic tumors. We employed Balb/c mice with bilateral breast cancer cell line 4T1 implants and randomized them into eight treatment groups: (i) Control, (ii) PP@T, (iii) anti‐PD‐1, (iv) PP@T+anti‐PD‐1, (v) PP@T+LIFU, (vi) PP@T+LIFU+anti‐PD‐1. The experimental procedure is outlined in Figure  6a. As shown in Figure 6b,c, PP@T‐augmented SDT effectively suppressed primary tumor growth but did not significantly affect distant tumors. Treatment with either anti‐PD‐1 or PP@T alone exhibited only a marginal effect on inhibiting both primary and distant tumors. Combination therapy with PP@T and anti‐PD‐1 resulted in a slight reduction in tumor growth, which did not reach statistical significance compared to single‐antibody treatments. However, the combination of SDT with anti‐PD‐1 significantly reduced tumor growth compared to the Control and single‐antibody treatment groups. Notably, within the combination groups, the PP@T+LIFU+anti‐PD‐1 group exhibited the most significant suppression of both primary and distant tumor growth and showed the greatest reduction in tumor weight. These results demonstrate that blockade with anti‐PD‐1 in combination with SDT and oxidative therapy yielded the most potent inhibitory effect on both primary and distant tumors.

Figure 6.

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a) Schematic illustration of the model and administration strategy for the treatment of primary and distant 4T1 tumor models. Curves representing initial tumor volumes b) and distant tumor volumes c) of mice treated with various treatments (n = 5). d) Schematic illustration of the therapeutic immunization on a lung metastasis mouse model. e) Weight of lung measured at the end of treatment. f) Inhibiting effect of various treatments on the formation of lung metastatic tumor. Statistical data were represented as the mean ± s.d. The level of statistical significance was indicated when appropriate (* P<0.05, ** P<0.01).

Inspired by the encouraging anti‐tumor efficacy of the PP@T in the bilaterally bearing 4T1 tumor model, we further assessed its anti‐metastatic performance using an aggressive whole‐body metastasis tumor model. As illustrated in Figure 6d, the experimental protocol involved the intravenous injection of additional 4T1 cells into orthotopic tumor‐bearing mice to induce lung metastasis. As shown in Figure 6e,f, the Control, PP@T, anti‐PD‐1, and PP@T + LIFU groups exhibited severe lung metastasis. However, the PP@T+ anti‐PD‐1 groups showed delayed aggressive lung metastasis, indicating that combining antibodies and oxidative therapy is crucial for inhibiting pulmonary metastasis. More importantly, the PP@T+LIFU+anti‐PD‐1 group showed nearly no metastasis, suggesting remarkable suppression of lung metastasis with our SDT in combination with the oxidative therapy and blockade strategy. Additionally, H&E staining of lung tissues confirmed metastasis in all groups except the PP@T+LIFU+anti‐PD‐1 group. Overall, PP@T‐amplified oxidative stress, in combination with blockade with anti‐PD‐1, exhibited robust ability to suppress lung metastasis.

2.8. Mechanism Study of Systematic Antitumor Immune Responses

To explore the underlying mechanism of the synergistic anti‐tumor effect induced by PP@T‐augmented oxidative stress in combination with PD‐1blockade, we examined immune cells within the tumors of mice using a bilaterally orthotopic model on day 11 after treatment. DC cells in lymph nodes play crucial roles in innate and adaptive immunities. According to Figures 7a and S22 (Supporting Information), the mature DCs (CD11c+, CD80+, and CD86+) rate of PP@T+LIFU+anti‐PD‐1 reached ≈81%. In contrast to other groups, it clearly activates the immune system. CD8+ T cells in tumor tissue were responsible for killing tumor cells. As shown in Figure 7b and Figure S23 (Supporting Information), mice treated with primary tumors subjected to PBS, PP@T injection alone, or anti‐PD‐1 treatment alone failed to promote the infiltration of CD8+ T cells into distant tumors. Additionally, the largest percentage of activated tumor‐infiltrating CD8+ T cells was observed in the PP@T+LIFU+anti‐PD‐1 treatment, suggesting that PP@T could be utilized in conjunction with therapeutic immune checkpoint inhibitors to achieve a better outcome for preventing tumor metastasis.

Figure 7.

Figure 7

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Anticancer immune responses. The proportion of a) mature DCs in lymph nodes and b) CD8+ T cells in the tumor and after various treatments by flow cytometry. c–f) The cytokines IFN‐γ, IL‐6, IL‐12, and TNF‐α in serum from differently treated mice. g) Immunofluorescence staining of CD8+ T cells (green) and IFN‐γ (red) in distant tumor sections. (G1: Control, G2: PP@T, G3: anti‐PD‐1, G4: PP@T+anti‐PD‐1, G5: PP@T+LIFU, G6: PP@T+ LIFU+anti‐PD‐1). Scale bar = 50 µm, Statistical data were represented as the mean ± s.d (n = 3).The level of statistical significance was indicated when appropriate (* P<0.05, ** P<0.01).

In addition, we analyzed the amount of tumor‐infiltrating IFN‐γ‐producing CD8+ (IFN‐γ+CD8+) T cells, which can directly kill cancer cells.[ 28 ] As shown in Figure S24 (Supporting Information), PP@T+LIFU+anti‐PD‐1 treatment led to significantly increased production of IFN‐γ+CD8+ T cells in distant tumors. Immunofluorescence assays further confirmed the anti‐tumor immunity triggered by PP@T. Visual examination revealed a more significant increase in CD8+ T cells and IFN‐γ+ CD8+ T cells following treatment with PP@T+LIFU+anti‐PD‐1 (Figure 7g). Additionally, we assessed the levels of immunosuppressive regulatory T cells (Tregs, CD3+CD4+Foxp3+) in tumors from the six experimental groups. PP@T+anti‐PD‐1 +LIFU significantly reduced the percentage of Tregs in distant tumors (Figure S25, Supporting Information). The collected serum of mice was subjected to ELISA assays to assess changes in multiple cytokines, including IFN‐γ, IL‐6, IL‐12, and TNF‐α. Similarly, as shown in Figure 7c‐f, PP@T+LIFU+anti‐PD‐1 resulted in a higher secretion level of these pro‐inflammatory cytokines.

Taken together, these findings establish a consistent pattern favoring an enhanced immune profile with the combination of PP@T‐amplified oxidative stress and anti‐PD‐1blockade increased CD8+T cell levels, reduced Foxp3+ Treg abundance, and heightened production of pro‐inflammatory cytokines such as IFN‐γ, IL‐6, IL‐12, and TNF‐α. These findings underscore the potential of the PP@T as an oxidative stress nano‐amplifiers to further amplify effector T cells, thereby triggering robust protective antitumor immunity capable of restraining tumor growth and metastasis.

3. Conclusion

In summary, our study emphasizes the possibility of using multifunctional nanosystem (PP@T) for amplifying oxidative stress and imaging‐guided oxidative/sonodynamic combination therapy in cancer. First, our in vitro and in vivo experiments illustrate that PP@T generates ROS, causing oxidative stress under the irradiation of LIFU. Moreover, released TH588 inhibits MTH1 protein activity and DNA repair processes, accumulating oxidative damage. Second, this combination therapy stimulates effective synergistic immunotherapeutic responses that inhibit primary tumor growth and metastasis. Our study demonstrate that TH588 and SDT can effectively amplify tumor oxidative stress in both living cells and in vivo, and more importantly, PP@T combined with anti‐PD‐1 blocking can produce a powerful anti‐tumor immune response, induced robust TILs recruiting to tumor bed, significantly improved the inhibition ability to primary tumor as well as antimetastasis. To the best of our knowledge, our oxidative stress nano‐amplifier were confirmed to be an effective synergistic therapy strategy and have significant clinical implications for developing new anti‐tumor strategies.

4. Experimental Section

Materials

Tetrakis (4‐carboxyphenyl) porphyrin (TCPP) was obtained from Tokyo Chemical Industry CO., Ltd. N,N’‐dimethylformamide (DMF), benzoic acid (BA) and Chloride octahydrate (ZrOCl2·8H2O) were purchased from Adama‐Beta Co., Ltd. Dopamine hydrochloride was obtained from Aladdin. Cell Counting Kit‐8, calcein‐AM, PI, and 2′,7′‐Dichlorofluorescin diacetate (DCFH‐DA) were purchased from Dojindo (Japan). TH588 were purchased from Selleck. All cell culture‐relevant reagents were obtained from the Beyotime Institute of Biotechnology (China). All commercial chemicals were used as received without further purification unless specified otherwise.

Synthesis of PCN‐224 and PP@T

PCN‐224 was synthesized according to a previously documented method.[ 18 , 19 ] In brief, 100 mg of TCPP, 300 mg of ZrOCl2•8H2O, and 2800 mg of benzoic acid were dissolved in 100 mL of DMF, respectively. Then, the mixture was stirred for 5 h at 90 °C in the dark. Finally, the resulting PCN‐224 was collected by centrifugation (10 000 rpm, 10 min) and washed with DMF and ethanol. For TH588 loading, the PCN‐224 (5 mg) and TH588 (4.5 mg) were dissolved in DMSO; then the mixture was stirred for 5 h. Finally, the product was washed with ethanol and deionized water. All supernatants were collected, and the drug load of TH588 was measured by HPLC.

To prepare PP@T, 10 mg of PCN‐224 @TH588 was dispersed in an ethanol‐water solution (15 mL, 3:4, v/v). 5 mg of Dopamine was injected into the above mixture solution under stirring. Subsequently, 20 mL of Tris(hydroxymethyl)aminomethane (10 mm, pH 8.5) was injected into the mixture and reacted for 48 h at room temperature in the dark. Finally, PP@T was collected by centrifugal washing 3 times with ethanol and water alternately. The preparation method of PCN‐224@PDA (PP) was the same as above, except that PCN‐224@TH588 was replaced by PCN‐224.

Detection of 1 O2

For the detection of 1 O2 by employing DPBF, an ethanol solution containing DPBF (10 mm) and PP@T (50 µg mL−1) was exposed to low‐intensity focused ultrasound (LIFU) irradiation (2.0 W cm−2) for different time durations. The absorption spectra were recorded every 2 min. To detect 1 O2 using ESR measurements, TEMP solution (20 µL) was mixed with PP@T dispersion (100 µg L−1), and the mixture was irradiated with US for different durations. The signal was recorded by an ESR spectrometer.

Cellular Experiments—Transmission Electron Microscopy (TEM) Assay

4T1 cells were seeded in 6‐well culture plates. After 12 h of incubation, cells were recovered for the uptake experiment. For apoptosis experiments, cells were incubated with a medium containing the PP@T nanoparticles at 50 µg mL−1 for 4 h. The cells in SDT groups were irradiated with US for 30 s. Cells were added to glutaraldehyde solution after being harvested, and TEM images were captured using a Hitachi TEM system.

Determination of Intracellular ROS

Intracellular ROS levels were measured using fluorescence microscopy and flow cytometry with DCFH‐DA, as the protocol reported previously. Briefly, 4T1 cells were incubated with a medium containing the PP@T nanoparticles at 50 µg mL−1 for 4 h. Then 10 µM of DCFH‐DA were added after washed with PBS. After treatment with LIFU for 30 s, cells were washed with PBS and imaged by microscope.

Cytotoxicity and Apoptosis Assay

4T1 cells were seeded in 96 well plates at a density of 1 × 103 cells mL−1 and in 35 mm cell culture dishes at a density of 1 × 105 cells mL−1 for 12 h. After incubation with different nanoparticles (PP, PP@T) for another 4 h, the cells in US groups were irradiated with LIFU for 30 sec. The cytotoxicity was determined by CCK‐8 assay or Calcein‐AM/PI staining assay. Apoptosis was evaluated by ANXA5/annexin V‐FITC and PI and detected by flow cytometry.

In addition, the mitochondrial membrane potential was estimated by using the JC‐1 staining assay. In brief, cells in different groups were treated with pre‐configured JC‐1 staining solution at 37 °C for 30 min. After the cells were digested and collected, the changes in mitochondrial membrane potential of the cells in different treatment groups were detected and analyzed by flow cytometry.

Colony Formation Assay

4T1 cells were seeded into 6 well plates at a density of 1 × 103 cells per well for 12 h. Then, cells were incubated with different nanoparticles for 4 h. The US groups were irradiated with LIFU for 30 s. The cells were incubated for 7 days with media change after every 2 days. Cells were fixed with 4% buffered formaldehyde for 30 min after 7 days. Colonies were dyed with 0.05% crystal violet solution for 20 min, then washed three times with PBS.

Fluorescence Imaging of DNA Mutant

Based on the high binding affinity of avidin to 8‐oxo‐dG,[ 29 ] the binding of 8‐oxo‐dG in DNA was imaged by Alexa Fluor 488 coupled avidin protein. 4T1 cells were inoculated in confocal dishes at a density of 1 × 105 per well. After incubation for 12 h, the medium was removed, and 1 mL pure medium, PP and PP@T Nps (50 µg mL−1) were added to each group for further incubation for 4 h. Then 4T1 cells were divided into the following groups: blank control group, LIFU group, PP@T group, PP + LIFU group, PP@T+ LIFU group. For the ultrasound irradiation group, cells were irradiated with low‐intensity focused ultrasound for 30 s (pulse mode, duty ratio 50%, 2 W cm−2) and continued to be incubated for 3 h. Then, cells were fixed in 4% paraformaldehyde at room temperature for 30 min and washed twice with TBST (Tris‐buffered saline containing 0.1% Triton X‐100). TBST solution containing 15% FBS was closed at room temperature for 2 h, and cells were incubated in TBST solution containing Alexa Fluor 488 conjugated avidin protein (10 µg mL−1) at 37 °C for another 1 h. The cells were then washed with TBST twice and stained with DAPI. Cells were observed under a confocal laser microscope, and the data were analyzed by Image J software.

Animal Experiments

Female Balb/c mice (6‐8 weeks old) weighted 18–19 g were purchased from Chongqing Medical University Laboratory Animal Center and kept in a SPF‐level sterile animal room. All live animal experiments were conducted in accordance with the criterion of the Animal Ethics Committee of Chongqing Medical University (No. (2021) 285). The living tumor model was established by injecting 4T1 cells suspended in 200 µL of aseptic PBS into the mammary fat pads on the right/left side of mice.

Fluorescence Imaging In Vivo

Tumor‐bearing mice were injected 200 µL (10 mg kg−1) of DIR‐labeled PP@T nanoparticles (10 mg kg−1) through the tail vein. IVIS imaging system was performed on the mice before injection and 6, 12, 24, and 48 h after the injection to detect the fluorescence intensity of DIR at different time points. The mice were sacrificed 48 h later, and the main organs and tumors were harvested for tissue imaging.

PA Imaging In Vivo

When the tumor volume reached 200 mm3, tumor‐bearing mice were injected with 200 µL (10 mg kg−1) of PP@T nanoparticles via the tail vein. PA imaging of tumors in mice was acquired after different intervals followed by PP@T administration.

In vivo Antitumor Studies

For in vivo treatment, the tumor‐bearing mice (tumor size reached 70–80 mm3) were randomly divided into 5 groups (n = 5): PBS group, LIFU group, PP@T group, PP + LIFU group, PP@T + LIFU. These mice were injected intravenously with PP@T or PP at the same TCPP dose (5 mg kg−1). In the US irradiation groups, mice were exposed to Ultrasound treatment (50% duty cycle, 2 W cm2, 5 min) 24 h post‐injection. The tumor size and weight of mice were recorded every other day. On day 14, mice were killed, and tumors and main organs were collected, dissected, and subjected to H&E, TUNEL, and PCNA immunofluorescence staining.

SDT Plus PD‐1 Blockade for Suppressing Distant Tumors

For the anticancer experiment, a bilateral tumor model was established. First, 4T1 cells were injected into the breast pad of each mouse as the primary tumor. After 7 days, the other breast pad was injected with 4T1 cells as the distant tumor (a simulation of metastatic tumor). Then mice were randomized to receive control (IgG 10 mg kg−1), anti‐PD‐1 (10 mg kg−1), PP@T, PP@T + anti‐PD‐1, PP@T +LIFU, PP@T + anti‐PD‐1 +LIFU. These mice were injected with PP@T intravenously (10 mg kg−1), and the primary tumors of mice were exposed to LIFU treatment (50% duty cycle, 2 W cm2, 5 min, one irradiation per three days, total 3 irradiations) 24 h post‐injection. Antibodies were administered i.p. on days 1, 4, 7 and 10. The tumor volume and weight of mice were recorded every other day. On the 14th day, mice were killed, and tumors and main organs were collected, dissected, and subjected to H&E, TUNEL, and PCNA immunofluorescence staining.

To establish a pulmonary metastasis model, 4T1 cells were injected intravenously via tail vein infusion. The following treatments were the same as the procedures above. Lungs were collected, fixed in Bouin's solution, and imaged at the end of the experiment.

Mechanism Investigation of the Combined Treatment

To systematically study the in vivo anti‐tumor immune response against simulated distant tumors, tumors, draining lymph nodes were collected, and single‐cell suspensions were prepared. The collected cells were further stained with several fluorescence‐bound antibodies and analyzed by flow cytometry. 1) CD8+ T cells (CD3+CD4−CD8+), 2) Tregs (CD3+CD4+Foxp3+), 3) DCs (CD11c+CD80+CD86+). Data were analyzed using FlowJo Analysis software. The intratumoral secretion levels of TNF‐α, IL‐6, IL‐12, and IFN‐γ were analyzed with ELISA kits according to the manufacturer's instructions.

Statistical Analysis

The data were expressed as mean ± s.d using GraphPad (Prism 6.0). One‐way analysis of variance statistical was used to calculate the experimental data. The data were categorized based on the p‐values and represented as “*” for p < 0.05 and “**” for p < 0.01.

Ethical Approval Statement

All animal experiments were approved by the Animal Ethics Committee of Chongqing Medical University.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

W.Z. and Y.R. contributed equally in this investigation. W.Z. performed conceptualization, methodology, formal analysis, funding acquisition, and Wrote original Draft. Y.R. performed conceptualization, methodology, formal analysis, and investigation. M.Y. performed methodology and formal analysis. Y.H. performed methodology and formal analysis. Z.W. performed methodology and resources. Y.C. performed supervision, H.R. performed conceptualization, methodology, formal analysis, validation, supervision, project administration, funding acquisition, and wrote, reviewed, and edited. All authors read and approved the final manuscript.

Supporting information

Supporting Information

ADHM-13-0-s001.docx (3MB, docx)

Acknowledgements

This work was supported by the National Natural Science Foundation of China (82071926), the Science and Technology Research Program of Chongqing Municipal Education Commission (Grant No. KJZD‐K202300403), the Natural Science Foundation of Chongqing (CSTB2023NSCQ‐MSX0149) and the CQMU Program for Youth Innovation in Future Medicine (No. W0170).

Zhang W., Ran Y., Yang M., Hu Y., Wang Z., Cao Y., Ran H., An Oxidative Stress Nano‐Amplifier for Improved Tumor Elimination and Combined Immunotherapy. Adv. Healthcare Mater. 2024, 13, 2402349. 10.1002/adhm.202402349

Data Availability Statement

The data generated in this study are available within the article or from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Information

ADHM-13-0-s001.docx (3MB, docx)

Data Availability Statement

The data generated in this study are available within the article or from the corresponding author upon reasonable request.

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