Metal-organic framework nanoparticles activate cGAS-STING pathway to improve radiotherapy sensitivity

Abstract

Tumor immunotherapy aims to harness the immune system to identify and eliminate cancer cells. However, its full potential is hindered by the immunosuppressive nature of tumors. Radiotherapy remains a key treatment modality for local tumor control and immunomodulation within the tumor microenvironment. Yet, the efficacy of radiotherapy is often limited by tumor radiosensitivity, and traditional radiosensitizers have shown limited effectiveness in hepatocellular carcinoma (HCC). To address these challenges, we developed a novel multifunctional nanoparticle system, ZIF-8@MnCO@DOX (ZMD), designed to enhance drug delivery to tumor tissues. In the tumor microenvironment, Zn²⁺ and Mn²⁺ ions released from ZMD participate in a Fenton-like reaction, generating reactive oxygen species (ROS) that promote tumor cell death and improve radiosensitivity. Additionally, the release of doxorubicin (DOX)-an anthracycline chemotherapeutic agent-induces DNA damage and apoptosis in cancer cells. The combined action of metal ions and double-stranded DNA (dsDNA) from damaged tumor cells synergistically activates the cyclic GMP-AMP synthase (cGAS)–stimulator of interferon genes (STING) pathway, thereby initiating a robust anti-tumor immune response. Both in vitro and in vivo experiments demonstrated that ZMD effectively activates the cGAS-STING pathway, promotes anti-tumor immune responses, and exerts a potent tumor-killing effect in combination with radiotherapy, leading to regression of both primary tumors and distant metastases. Our work provides a straightforward, safe, and effective strategy for combining immunotherapy with radiotherapy to treat advanced cancer.

Graphical Abstract

Introduction

Liver cancer is one of the most common cancers in the world, with high morbidity and mortality rates, seriously affecting patients’ quality of life, and hepatocellular carcinoma (HCC) is the most common type of liver cancer [1, 2]. Cancer immunotherapies have changed the paradigm of cancer treatment; these therapies are designed to activate or boost the activation of the immune system, inhibit the immune escape of tumor cells, and have fewer off-target effects than chemotherapy and other drugs that directly kill cancer cells [3,4,5,6]. However, due to the complex microenvironment of the liver (chronic liver disease background) and HCCs, immune checkpoint inhibitor (ICI) immunotherapy still faces great challenges, such as low response rates, drug resistance, and potential systemic side effects. New precision immunotherapies are therefore urgently needed [7]. As an innate immune signaling pathway that recognizes cytoplasmic DNA accumulation arising from multiple perturbations of cellular homeostasis and triggers an effective anti-tumor immune response, cyclic GMP-AMP synthase (cGAS)–stimulator of interferon genes (STING) activation therapies is receiving increasing attention [8,9,10]. However, commercially available cGAS-STING agonists are soluble and easily cleared. Furthermore, soluble agonists elicit the broad cytokine responses that may lead to uncontrolled inflammation and fatal injury [11]. Therefore, the development of new cGAS-STING activators with high targeting and low toxicity is imperative.

Radiotherapy is one of the major cancer treatments that promotes local tumor cell killing and induces tumor microenvironment (TME) immunomodulation [12, 13]. Radiotherapy-induced DNA damage response is critical for tumor cell killing by radiotherapy, and DNA double-strand breaks (DSBs) further stimulate activation of the cGAS-STING pathway [14]. Therefore, radiotherapy combined with cGAS-STING pathway activators is promising. However, tumor radiosensitivity is a major obstacle limiting the efficacy of radiotherapy, and conventional radiosensitizers appear to be unsatisfactory for HCC [15]. Modulation of oxidative stress in cancer cells is increasingly becoming a popular anticancer strategy, and there is growing evidence that inhibitors of the antioxidant system can promote tumor cell radiosensitivity [16, 17]. The Fenton or Fenton-like reaction that occurs when metal ions react with hydrogen peroxide (H₂O₂) in the TME generates cytotoxic hydroxyl radicals (-OH), which induces cellular oxidative stress and increases the sensitivity of tumor cells to radiotherapy [18, 19]. Therefore, it is theoretically feasible to deliver metal ions in a safe manner to exert antitumor effects and enhance the sensitivity of tumor cells to radiotherapy. Chemotherapeutic agents for radiation therapy sensitization have been widely used in the clinic. Some chemotherapeutic agents, such as doxorubicin (DOX), can also induce immunogenic cell death (ICD) and consolidate radiotherapy-induced antitumor immunity. However, their therapeutic efficacy is limited due to drawbacks such as instability, rapid elimination and non-specific tumor homing ability, more stable and effective delivery vehicles are needed [20,21,22].

Zeolite imidazole framework 8 (ZIF-8) is an emerging class of metal-organic frameworks (MOFs) with properties such as simple synthesis, easy modification, high encapsulation rate, and pH-responsive degradation [23, 24]. ZIF-8 protects the drug from premature release, improves drug enrichment at the tumor site, and reduces systemic toxicity. Zn²⁺, a decomposition product of ZIF-8 in the acidic environment of tumors, generates reactive oxygen species (ROS) through the Fenton-like reaction, which increases the sensitivity of tumor cells to radiotherapy [25]. In addition, Zn²⁺ can promote the phase separation of cGAS-DNA, enhance the activity of cGAS enzyme, and promote anti-tumor immune response [24, 26]. Therefore, we chose ZIF-8 as the drug carrier to design the nanoparticles (NPs).

Based on the above background, we designed a therapeutic strategy using ZIF-8 loaded with MnCO and DOX to form multifunctional NPs (ZIF-8@MnCO@DOX, ZMD) (Fig. 1). ZIF-8 protects the drug from clearance and releases DOX, Mn²⁺, and Zn²⁺ in the acidic environment of the tumor. Mn²⁺ and Zn²⁺ undergo a Fenton-like reaction that promotes ROS production and apoptosis in tumor cells and enhances cellular radiosensitivity. CO further induces apoptosis in cancer cells by reducing cellular protein synthesis through inhibition of cellular mitochondrial respiration [27, 28]. DOX, an antibiotic analog, induces DNA damage and apoptosis in tumor cells. The dsDNA released from tumor cell damage activates the cGAS-STING pathway, and Zn²⁺ promotes cGAS-DNA phase separation to further enhance cGAS enzyme activity and induce a strong anti-tumor immune response [29]. Our work aims to provide a simple, safe, and effective approach for immunotherapy combined with radiotherapy in the treatment of advanced cancer.

Scheme 1

Schematic illustration of ZIF-8@MnCO@DOX (ZMD NPs) for cancer therapy. ZMD NPs are synthesized by an in situ “one-pot” self-assembly method. ZIF-8 protects the drug from being released and scavenged in neutral environments, whereas it releases Zn²⁺, Mn²⁺, CO and DOX in the low pH and high H₂O₂ level environment of tumors. Mn²⁺ and Zn²⁺ undergo a Fenton-like reaction that promotes ROS production, thereby killing tumor cells and enhancing cellular sensitivity to radiotherapy. Additionally, DOX is a chemotherapeutic agent that kills tumor cells. The dsDNA released from cellular damage together with metal ions Zn²⁺ and Mn²⁺ activate the cGAS-STING pathway, inducing a strong anti-tumor immune response. Created with Biorender. com

Materials and methods

Materials

Zn (NO₃)₂-6H₂O was purchased from Innochem (Peking, China), 2-Methylimidazole, MnCO were purchased from Aladdin’s Reagent (Shanghai, China). DOX was provided by Yuan Ye (Shanghai, China). Calcein AM, DNA damage detection kit, ROS assay kit, and crystal violet reagent were purchased from Beyotime Biotechnology (Shanghai, China). CCK-8 cell proliferation and cytotoxicity assay kit was provided by Solarbio (Peking, China). Apoptosis detection kit was purchased from Vazyme (Nanjing, China). Matrigel and transwell plates were supplied by Corning, USA. 4% paraformaldehyde was provided by Biosharp (Peking, China). FITC fluorescent dye was purchased from MedChemExpress (Shanghai, China). Dulbecco’s modified eagle medium (DMEM) and fetal bovine serum (FBS) were purchased from Thermo Fisher Scientific (USA). Rabbit polyclonal antibody to STING (DF12090), rabbit polyclonal antibody to phospho-STING (AF7416), rabbit polyclonal antibody to phospho-IRF3 (AF2436) and rabbit polyclonal antibody to phospho-TBK1 (AF8190) were purchased from Affinit (Jiangsu, China). β-Actin mouse antibody, anti-rabbit or anti-mouse horseradish peroxidase (HRP)-labeled secondary antibodies were purchased from Abmart (Shanghai, China). Anti-CD11c antibody (E-AB-F0991C), anti-CD86 antibody (E-AB-F0994D), anti-CD80 antibody (E-CK-A107), anti-CD8 antibody (E-AB-F1104D), anti-CD4 antibody (E-AB-F1097C) were purchased from Elabscience (Shanghai, China). Anti-CD45 antibody (557659), anti-F4/80 antibody (565411), anti-CD11b antibody (552850), and anti-CD206 (568808) antibody were purchased from BD Biosciences (USA). Primers for qPCR were purchased from Tsingke (Peking, China). Primer sequences are shown in Supplementary Table S1.

Preparation of ZMD NPs

Synthesis of ZIF-8

92 mg of dimethylimidazole (2-MIM) was dissolved in 10 ml of methanol, and subjected to sonication for 2 min to achieve complete dissolution. 43.2 mg of Zn (NO₃)₂-6H₂O was dissolved in 10 ml of methanol, and sonicated for 2 min until fully dissolved. The two solutions were mixed and stirred at room temperature at a speed of 1,000 rpm for 1 h. A white suspension was obtained after the reaction. The synthesized NPs were collected by centrifugation at 12,000 rpm for 10 min, washed three times with methanol, and freeze-dried to obtain ZIF-8 NPs (Scheme 1).

Synthesis of ZIF-8@MnCO

ZIF-8@MnCO was prepared by dissolving 10 mg of MnCO in 10 ml of methanol, 92 mg of 2-MIM in 10 ml of methanol, 43.2 mg of Zn (NO₃)₂-6H₂O in 10 ml of methanol, and each solution is sonicated for 2 min to ensure complete dissolution. The MnCO solution was then added to the Zn (NO₃)₂-6H₂O solution and gently mixed. Following this, the combined solution was added dropwise to the 2-MIM solution at a rate of one drop every 5 s, while continuously stirring at a speed of 1000 rpm. The reaction was allowed to proceed at room temperature for 1 h. The resulting products were collected by centrifugation (12,000 rpm,10 min), washed three times with methanol, and stored after freeze-drying.

Synthesis of ZIF-8@MnCO@DOX

The synthesis of ZIF-8@MnCO@DOX was executed using an in situ ‘one-pot’ approach. Initially, 10 mg of MnCO was dissolved in 10 ml of methanol, 92 mg of 2-MIM was dissolved in 10 ml of methanol, and 10 mg of DOX was dissolved in 10 ml of distilled water, with each solution sonicated for 2 min to ensure complete dissolution. Next, 43.2 mg of Zn (NO₃)₂-6H₂O was dissolved in the DOX solution. The MnCO solution was then incorporated into the mixture of DOX and Zn (NO₃)₂-6H₂O, followed by gentle shaking to ensure uniform mixing. The mixed solution was added dropwise into the 2-MIM solution utilizing a dropping funnel, at a rate of one drop every 5 s, while continuously stirring with a speed of 1,000 rpm. Once the dripping was completed, the reaction was covered with plastic wrap and stirred at room temperature for an additional hour. The product was collected via centrifugation (12,000 rpm,10 min), washed three times with distilled water and freeze-dried to prepare ZMD.

Characterization of ZIF-8@MnCO@DOX

The size and zeta potential of the NPs were measured by dynamic light scattering (DLS). The morphology of the NPs was observed by transmission electron microscopy (TEM). Elemental mapping was used to confirm the elemental composition of the NPs.

The DOX absorption peak was detected by UV spectrophotometry and the standard curve of DOX was plotted as a basis for the calculation of DOX concentration. The release of DOX from ZMD was studied by dialysis assay. A certain amount of ZMD was filled into a dialysis bag and incubated in solvents of different pH at room temperature with stirring. 1 ml of solvent was removed and replenished with fresh solvent at different time points, followed by measurement of DOX in the solvent using a fluorescence spectrophotometer. The formula for calculating the amount of DOX released is as follows:

$$\:Cumulative\:release\:amount=\frac{{M}_{t}}{{M}_{total}}\times\:100\%$$

M_t is the amount of DOX released into the solvent at the time point, M_total is the total amount of DOX in the dialysis bag.

To evaluate the release of manganese ions from ZMD, a quantity of ZMD was placed in phosphate buffer solution (PBS). The filtrate after ultrafiltration was collected at predetermined time points, the release of manganese ions was detected by Inductively Coupled Plasma (ICP), and the cumulative release was calculated.

Stability testing of ZMD

An amount of ZMD was dissolved in DMEM medium containing 10% FBS and PBS buffer with pH 7.4 (n = 3), respectively. The particle size change of ZMD in the sample solution was measured at intervals.

Cell culture

Human HCC cell huh7 and mouse HCC cell hepa1-6 were purchased from Procell Life Science & Technology Co., Ltd. (Wuhan, China). The cell lines were cultured in DMEM supplemented with 10% FBS and 1% penicillin-streptomycin liquid. All cells were cultured in a humidified incubator at 37 °C in the presence of 5% CO₂.

Cytotoxicity assay in vitro

HCC cells were seeded in 96-well plates at a density of 3000 cells per well. Each well was treated by adding different concentrations of different agents for 24 h. After the cells were attached to the wall, the old medium was aspirated and 100 µl of medium containing 10 µL of CCK-8 was added, and the plates were incubated at 37 °C for 2 h in a light- protected environment. After that, the absorbance of each well was measured at 450 nm wavelength and the half maximal inhibitory concentration (IC50) value was calculated.

Cellular uptake assay

FITC dye was incubated with ZMD to synthesize fluorescently labelled ZMD. FITC-ZMD was incubated with HCC cells for 4 h. After that, the medium was aspirated and the cells were washed three times with PBS to remove the drug that did not enter the cells. The cells were stained with 2-(4-amidinophenyl)-6-indolylamidine dihydrochloride (DAPI) and imaged using a laser confocal microscope. The experiment was repeated and cells were collected to detect FITC fluorescence intensity in the cells using a flow cytometer (CytoFlex, Beckman Coulter).

Live-dead staining assay

HCC cells in logarithmic growth phase were inoculated into six-well plates at a density of 2 × 10⁵ cells per well and cultured for 8 h in fresh medium containing 30 ug/ml of different NPs before being subjected to 6 Gy radiation treatment. After 24 h of incubation, 1 mL of live-dead staining solution was added, and the cells were incubated in an incubator protected from light for 15 min. The cells were observed and photographed using a fluorescence inverted microscope at 490 nm and 533 nm excitation wavelengths.

Wound healing assay

Cells were seeded in 6-well plates and scraped vertically with a 200 µl pipette when the cells reached approximately 90% confluency. After washing three times with PBS, images of 0 h HCC cells were taken. Next, the cells were treated with different NPs and underwent radiotherapy, and continued to be cultured with medium containing 2% FBS for 24 h. The images were captured with the microscope.

Transwell invasion assay

Cells from different treatment groups were pre-incubated with serum-free DMEM medium for 12 h, and then inoculated into the upper chamber of transwell plates (pre-coated with 70 µL of Matrigel in the upper chamber of the transwell plates and incubated at 37℃ for 4 h), and the lower chamber was added with DMEM containing 20% of FBS. After 24 h of incubation, the infiltrating cells were fixed with formalin, and stained with crystal violet for 15 min. and then observed and counted by inverted microscope (Olympus Corporation).

Cell colony formation assay

HCC cells were inoculated in 6-well plates at a density of 5 × 10³ cells/well and cultured for 12 h. Then the cells were treated with medium containing different agents (at a concentration of 30 µg/ml) for 8 h and the radiotherapy group was irradiated with 6 Gy. The cells were continued to be cultured for 1 week. Cells were fixed with paraformaldehyde and stained with crystal violet reagent. The number of infiltrating cells was observed and counted using an inverted microscope (Olympus).

Detection of intracellular ROS

HCC cells were inoculated into six-well plates at a concentration of 2 × 10⁵ cells/well. After the cells were wall-adhered, they were treated with different NPs for 8 h. After fluid exchange, the radiotherapy group was irradiated with 6 Gy, and the culture medium was removed after 24 h of further incubation. The cells were washed three times with PBS, then treated with 5 µM DCFH-DA solution, and incubated at 37 °C for 30 min in light-avoidance incubation. The amount of ROS production was detected by flow cytometry.

γH2Ax fluorescence detection

2.0 × 10⁵ cells per well were inoculated into six-well plates and cultured for 24 h. Then the medium was replaced with new medium containing different NPs (concentration of 30 µg/mL) for 8 h. Subsequently, the cells in the radiotherapy group were irradiated with 6 Gy and continued to be cultured for 24 h. After cell fixation, cells were incubated with γH2Ax primary antibody for 1 h at room temperature, followed by goat anti-rabbit IgG labeled primary antibody. Imaging analysis was performed under a microscope.

Western blot

Cell samples were lysed on ice in radio immunoprecipitation assay (RIPA) lysis buffer containing protease inhibitors and phosphatase inhibitors. After centrifugation at 12,000 rpm for 20 min, the supernatant of each sample was collected and the protein concentration in the samples was determined by the bicinchoninic acid assay (BCA). All protein samples were stored at -80 or -20 °C. Proteins in each sample were separated using a 10% sodium dodecyl sulfate (SDS) polyacrylamide gel at 80 V and then transferred to polyvinylidene fluoride (PVDF) membranes (Millipore, NJ, USA) at 200 mA. The membranes were blocked with 5% skimmed milk for 1 h at room temperature and then incubated overnight at 4 °C with the corresponding primary antibody. Anti-rabbit or anti-mouse HRP-labeled secondary antibodies were incubated for 1 h at room temperature, and the bands were visualized with a chemiluminescent kit.

Apoptosis detection

Cells were collected by trypsinization (without ethylenediamine tetra acetic acid) and washed twice with PBS, then stained with Annexin V and propidium iodide (PI) at a concentration of 50 µg/mL for 15 min at room temperature in a dark room. The apoptosis level was detected by flow cytometry and analyzed by FlowJo.

Maturation and activation of bone marrow-derived dendritic cells (BMDCs)

Mouse femoral and tibial bone marrow cells were cultured with RIPM 1640 medium containing GM-CSF (1000 U/ml) and IL-4 (1000 U/ml). The medium was changed every other day and the cells were collected on day 7 for subsequent studies. Dendritic cell (DC) cells treated with different drugs were collected, washed with PBS and stained with anti-CD80, anti-CD86 and anti-CD11c antibodies. The maturation of BMDC cells was then detected by flow cytometry.

Reverse transcription quantitative polymerase chain reaction (RT-qPCR)

RNA was extracted from cells using Trizol reagent and RNA purification kit, and the quantity and quality of RNA was confirmed using a NanoDrop spectrophotometer. Next, we synthesized cDNA using a reverse transcription kit with gDNA remover (Vazyme). 2 × Color SYBR Green qPCR Master Mix (Vazyme) was used for RT-qPCR experiments on the light cycler 480 instrument (Roche).

RNA sequencing (RNA-seq)

Absolute quantitative transcriptome sequencing was entrusted to the Institute of Hydrobiology, Chinese Academy of Sciences. 2.0 × 10⁵ cells per well were seeded and cultured in six-well plates for 24 h, then replaced with a new medium containing PBS or ZMD (concentration of 30 µg/mL) for 24 h. Cells were collected, mRNA was enriched from total RNA using oligo (dT) magnetic beads, and mRNA fragments were sheared into short fragments of approximately 300 bp using lysis buffer. The mRNA was used as a template to synthesize cDNA, and PCR amplification was performed after purification and modification to ensure that the effective concentration of the library was greater than 2 nM. The sequencing was performed on the Illumina platform. RNA sequencing results were analyzed using R version 4.2.0 (https://www.r-project.org/). Differential gene expression analysis was performed using the R package “DESeq2” with the thresholds of Fold Change ≥ 2 and FDR < 0.05. Heatmap was realized by “Pheatmap” R package. KEGG and GO pathway enrichment analysis was performed using the R package “ClusterProfler”, and GSEA analysis was conducted with GSEA software.

Immunofluorescence

After the cells or tumor tissues were fixed and punched, they were incubated with the corresponding primary antibody for 1 h at room temperature, then incubated with the corresponding anti-rabbit or anti-mouse fluorescently labelled secondary antibody for 50 min at room temperature before being imaged and analyzed under the microscope.

Tumor model establishment, in vivo fluorescence imaging and anti-cancer therapy

C57BL/6 mice (male, 4–6 weeks) were purchased from Wuhan sblbio Co. Ltd. and housed in a specific pathogen-free (SPF) facility. All animal experiments were conducted under the guidelines approved by the Experimental Ethics Committee of the First Clinical College of Wuhan University (Institutional Animal Care and Use Committee issue number: WDRM 20240906 C). 1 ml of PBS solution containing 7 × 10⁶ hepa1-6 cells was injected subcutaneously into the right lumbar rib side of C57 mice. When the tumor volume increased to 500 mm³, 100 µL of saline containing FITC-labelled NPs (1 mg/ml) was injected into the tail vein of the mice. The fluorescence signals on the tumors at different time points were measured using the IVIS imaging system while maintaining inhalation anesthesia. Twenty-four hours after drug injection, mice were euthanized, and tumor tissues and major organs were collected for fluorescence imaging to observe the distribution of the drug in the body.

After modelling as described above, tumor volume and mouse body weight were measured every other day. After three times of injections of NPs into the tail vein and three times of radiotherapy treatments (n = 5), the mice were euthanized, and orbital blood, tumor tissue, and major organs were collected for subsequent analysis.

In the lung metastasis model, mice were injected with 100 µl of saline containing 1 × 10⁶ hepa1-6 cells via tail vein to generate lung metastases. Subsequently, the mice were randomly divided into 8 groups (n = 3) to receive drug and radiation treatment. 30 days later, the mice were euthanized to obtain lung tissue sections for immunohistochemical experiments.

Flow cytometry analysis on tumor tissues

Tumor tissues were split into small pieces and incubated with medium containing 0.8 mg/mL collagenase D, 0.2 mg/mL DNA zyme, and 0.1 mg/mL hyaluronic acid (Meilunbio, China) at 37 °C for 30 min, and then single-cell suspensions were obtained by passing through 70 μm nylon mesh. After Percoll (Meilunbio, China) was added to the cell suspension, the cells in the lower layer were collected by gradient centrifugation and closed, labeled with immunocyte markers using appropriate antibodies, and incubated for 30 min at room temperature before being analyzed on a flow cytometer. Figure S9 shows the exact process.

Blood biochemical index

Blood collected from the orbits of each treatment group (n = 3) was centrifuged at 3000 rpm for 15 min at 4 °C to collect serum, which was analyzed on an Impact 400 clinical biochemical automatic analyzer (USA, Gilford) for ALT, AST, UA, CREA, and other biochemical parameters.

H&E staining

After all mice were sacrificed, lungs, hearts, spleens, livers and kidneys were collected and stained with hematoxylin-eosin (H&E) staining, and the pathology of each tissue was observed under a microscope.

Statistical analyses

The measurement data were expressed as the mean ± standard deviation (SD). All statistical analyses were performed using Prism software (GraphPad Prism version 10.1.2; www.graphpad.com). Statistical analysis was performed by two-tailed Student’s t-test for comparison between two groups and one-way analysis of variance (ANOVA) followed by Turkey’s post-test for comparison of three or more groups. Survival rates of mice were compared using the Kaplan-Meier method. p values < 0.05 was considered statistically significant; “ns” represented non-significance; *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Result

Synthesis and characterization of ZMD

We used an in situ ‘one-pot’ method to prepare ZMD. TEM shows successful synthesis of homogeneous NPs with a particle size of about 100 nm (Fig. 1A). DLS further verified that the particle size of ZMD is about 100 nm, which is consistent with the TEM results, and that the ZMD surface carries a positive charge of about 10 mv (Fig. 1B, C). In general, NPs smaller than 10 nm in diameter are rapidly cleared by the kidney, whereas NPs larger than 200 nm in diameter are at risk of activating the complement system. The size of 100 nm helps our NPs to extravasate and penetrate deeper into the tumor tissue through the enhanced permeability and retention effect (EPR) effect [30, 31]. Furthermore, it was demonstrated that NPs with a negative surface charge exhibited more affinity toward internalization than NPs with a negative surface charge [32]. The elemental mapping confirmed the presence of Mn, N, O and Zn elements in ZMD (Fig. 1D). Moreover, we examined the release of manganese ions and DOX from ZMD at different pH values. The results showed that under neutral conditions, the NPs could maintain stability for a long time and protect the drug from being released, while under acidic conditions, the release rate of manganese ions could be up to 95% in 30 h and DOX could be up to 82% in 8 h (Fig. 1E, F). To further demonstrate the stability of ZMD under neutral conditions, we measured the particle size change of ZMD in PBS and DMEM solution containing 10% FBS for 8 consecutive days. The results showed that ZMD could be stabilized in solution for a long period of time (Figure S1). After labeling ZMD with FITC, we observed the uptake of ZMD by hepa1-6 and huh7 cells using confocal microscopy. The results showed a high uptake of ZMD by HCC cells at 4 h of co-incubation (Fig. 1G). Flow cytometry further validated the uptake of ZMD by HCC cells and obtained results consistent with confocal microscopy (Fig. 1H, J, K). To investigate the toxicity of different NPs on HCC cells and the optimal drug concentration, we performed cell counting kit-8 (CCK8) experiments to calculate the IC50 of NPs. Figure 2L shows the IC50 values of various agents against hepa1-6 and huh7 cells. The results showed that in hepa1-6 cells, the IC50 values of ZIF-8, ZIF-8@MnCO and ZMD were 67.17 µg/ml, 52.69 µg/ml, and 30.71 µg/ml respectively. In huh7 cells, the IC50 values of ZIF-8, ZIF-8@MnCO, and ZMD were 59.19 µg/ml, 44.63 µg/ml and 28.71 µg/ml respectively. The results indicated that ZMD had the best killing effect on tumor cells.

Fig. 1

Preparation and characterization of ZMD. (A) TEM observation of the morphology of ZMD. DLS measured the particle size of ZMD to be about 100 nm (Scale bar = 100 nm) (B), and the charge of ZMD to be about + 10 mv (C). (D) Elemental scanning verified that ZMD contained the elements Mn, N, O and Zn (Scale bar = 100 nm). Release of Mn²⁺ (E) and DOX (F) with time at different pH values. (G) Confocal microscope shot of ZMD uptake by hepa1-6 and huh7 cells. Scale bar = 100 μm. (H) The uptake of ZMD by hepa1-6 and huh7 cells was detected by flow cytometry. (K) Quantitative analysis of FITC fluorescence intensity detected by flow cytometry. (L) IC50 values of ZIF-8, ZIF-8@MnCO, and ZMD acting on hepa1-6 and huh7 cells. ****p<0.0001

Tumor-killing effect of ZMD in vitro

To evaluate the role of ZMD in inhibiting tumor cell growth, invasion and metastasis, we performed wound healing, transwell and clone formation assays on hepa1-6 and huh7 cells. The wound healing assay confirmed that ZMD significantly inhibited the metastasis of HCC cells, and the combination of ZMD and radiotherapy could exert a powerful inhibitory effect on the metastasis of HCC cells (Fig. 2A, Figure S2B). Additionally, transwell assay showed that ZMD significantly inhibited the invasive ability of HCC cells compared with ZIF-8 and ZIF8@MnCO (Fig. 2B, Figure S2A). The results of clone formation assays showed that ZMD synergized with radiotherapy better inhibited the colony formation ability of HCC cells (Fig. 2C, Figure S2C). Moreover, live and dead cell staining assays further confirmed that ZMD significantly killed tumor cells, and the activity of HCC cells was significantly inhibited when treated by ZMD combined with radiotherapy (Fig. 2D, E).

We evaluated intracellular ROS generation after treatment with different NPs using a DCFH-DA assay kit. The results showed that ZMD synergized with radiotherapy significantly induced intracellular ROS generation. Compared with ZIF-8, ZIF-8@MnCO and DOX, ZMD induced ROS generation more significantly (Fig. 2F). Figure 2G and Figure S3 showed that ZMD could effectively induce apoptosis in tumor cells, and the apoptotic effect of HCC cells was more significant after being co-treated with ZMD and radiotherapy.

Fig. 2

Tumor cell killing ability of ZMD NPs. (A) Quantitative analysis of migration distance of hepa1-6 and huh7 cells after different NPs treatments and x-ray (6 Gy) irradiation (n = 3). (B) Quantitative analysis of the invading cell numbers of hepa1-6 and huh7 cells after different NPs treatments and x-ray (6 Gy) irradiation (n = 3). (C) Quantitative analysis of clone formation of hepa1-6 and huh7 cells after different NPs treatments and x-ray (6 Gy) irradiation (n = 3). (D) Quantitative analysis of the cell viability of hepa1-6 cells after different NPs treatments and x-ray (6 Gy) irradiation (n = 3). (E) The survival rate of hepa1-6 cells after treatment with different drugs and X-ray irradiation was characterized by live and death cell staining assays. Scale bar = 100 μm. n = 3. (F) Effect of different NPs treatments and x-ray (6 Gy) irradiation on intracellular ROS production. (G) Effect of different NPs treatments and x-ray (6 Gy) irradiation on cell apoptosis. n = 3. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

ZMD NPs activate the cGAS-STING pathway in HCC cells

dsDNA released into the cytoplasm by cellular injury triggers activation of the cGAS-STING pathway. dsDNA binds to cGAS and promotes the synthesis of 2′3′ cyclic GMP-AMP (cGAMP). cGAMP binds to STING dimer located in the endoplasmic reticulum (ER) membrane and promotes the recruitment of TANK-binding kinase 1 (TBK1) by STING, which facilitates the autophosphorylation of TBK1, phosphorylation of STING, and interferon regulatory factor 3 (IRF3) recruitment and phosphorylation, thereby inducing the expression of type I interferons (IFNs), interferon-stimulated genes (ISGs) and several other inflammatory mediators, pro-apoptotic genes and chemokines [26, 33]. Immunofluorescence results showed that ZMD caused DNA damage in HCC cells, and the combination of ZMD and radiotherapy produced a strong DNA damage effect (Fig. 3A, Figure S4). The immunofluorescence results verified that ZMD combined with radiotherapy potently activated the cGAS-STING pathway, and the activation effect of ZMD on cGAS-STING pathway was more pronounced compared to ZIF-8, ZIF-8@MnCO and DOX (Fig. 3B, C, D). Western blotting results further verified the induction of TBK1, IRF3 and STING phosphorylation by ZIF-8, ZM, DOX and ZMD. The activation of the cGAS-STING pathway was further amplified when the drugs were applied in combination with radiotherapy, and the combination of ZMD and radiotherapy strongly activated the cGAS-STING pathway (Fig. 3E, Figure S5A, B, E and F).

Fig. 3

ZMD activates the cGAS-STING pathway in hepa1-6 cells. (A) Effect of different NPs treatments and x-ray (6 Gy) irradiation on γ-H2AX protein production in hepa1-6 cells. n = 3. Scale bar = 50 μm. (B) Effects of different NPs treatments and X-ray irradiation (6 Gy) treatments on p-IRF3 and p-STING expression in hepa1-6 cells. n = 3. Scale bar = 50 μm. Quantitative analysis of the fluorescence intensity of p-IRF3 (C) and p-STING (D). n = 3. (E) Activation of cGAS-STING pathway in hepa1-6 cells by different NPs treatments and x-ray (6 Gy) irradiation. n = 3. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

ZMD NPs promote DC cells maturation and anti-tumor immune response

Activation of cGAS-STING pathway promotes the expression of IFNs. IFNs accomplish antigen presentation by stimulating DC cells and macrophages maturation and activation, increase granzymes and perforin production by cytotoxic T cells (CTL) and natural killer cells (NK cells), and activates innate and adaptive immunity to kill cancer cells [34]. Therefore, we treated DC cells with different drugs to explore the role of ZMD in promoting DC cell maturation and DC cell cytokine release. We found that ZMD significantly promoted DC cell maturation compared with ZIF-8, ZIF-8@MnCO and DOX (Fig. 4A, E). Figure 4B, C and D showed that ZMD significantly promoted the expression of IFNβ, CXCL10 and ISG15 in DC cells. The results indicated that ZMD could induce DC cells to release cytokines to perform anti-tumor immunotherapy. Additionally, the results of western blot showed that the cGAS-STING pathway was significantly activated in DC cells after treatment with ZMD (Fig. 4F).

We further explored the mechanism of anti-tumor effects of ZMD by transcriptome sequencing. We identified 232 genes that were differentially expressed in the PBS and ZMD-treated groups (Fig. 4G). The results showed that tumor suppressor genes such as Atoh8, Mt2, and Mt1 were up-regulated in the ZMD-treated group, and genes promoting HCC progression such as Ccn2, Ahnak2, and Map3k14 were down-regulated, suggesting that ZMD can exert anti-tumor effects (Fig. 4H). GO enrichment analysis revealed that the up-regulated genes in ZMD-treated group were related to the pathways of regulation of cell cycle, positive regulation of apoptotic process, negative regulation of transforming growth factor beta receptor signaling pathway and intrinsic apoptotic signaling pathway in response to DNA damage by p53 class mediator (Fig. 4I). KEGG enrichment analysis has shown that the deferentially expressed genes were related to the NF-kappa B signaling pathway (Fig. 4J). GSEA analysis revealed that ZMD was associated with the NF-kappa B signaling pathway, the regulation of B cell activation, the regulation of cell killing, the regulation of immune response to tumor cell, the regulation of T cell mediated cytotoxicity and T cell activation (Figure S6). These results suggested that ZMD treatment was associated with tumor- as well as immune-related pathways in HCC cells.

Fig. 4

ZMD NPs promotes the maturation of BMDCs and anti-tumor immune response. (A) Flow cytometry detection of DC cell maturation after different NPs treatments. n = 3. (B-D) Effects of different NPs treatments on IFNβ (B), CXCL10 (C), ISG15 (D) expression in DC cells. n = 3. (E) Quantitative analysis of DC cell maturation after different NPs treatments. n = 3. (F) Effects of different NPs treatments on cGAS-STING pathway activation in DC cells. n = 3. (G) Volcano plots of differentially expressed genes in ZMD- and PBS-treated hepa1-6 cells. (H) Heatmap presentation of differentially expressed genes in ZMD- and PBS-treated hepa1-6 cells. (I) GO enrichment analysis of highly expressed genes in ZMD-treated groups. (J) KEGG enrichment analysis of differentially expressed genes in ZMD-treated groups. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

Biocompatibility and therapeutic effect of ZMD in vivo

To further validate the safety and therapeutic efficacy of ZMD, we conducted in vivo studies in mice. We performed tail vein administration on days 7, 9, and 11 after subcutaneous tumor modelling and 6 Gy of radiotherapy on the day following the drug treatment, and euthanized the mice on day 14 (Fig. 5A). We collected blood from mice for blood biochemistry analysis and major organs for immunohistochemistry to investigate the compatibility of ZMD. The results showed that ZMD had no significant toxicity in mice. There were no obvious abnormalities in blood biochemistry (Figure S7), nor was there any significant damage to the heart, liver, spleen, lungs, and kidneys in the drug-treated mice (Figure S8). The safety of ZMD treatment was further confirmed by the fact that ZMD had no significant effect on the body weight of the mice during the treatment period (Fig. 5B). ZMD inhibited tumor growth in mice more significantly than ZIF-8 and ZIF-8@MnCO, and the combination of ZMD with radiotherapy produced therapeutic effects on tumors and significantly reduced tumor burden in mice (Fig. 5C, D, E). We performed animal in vivo imaging experiments to verify the tumor targeting of ZMD after tail vein injection at 0 h, 2 h, 4 h, 6 h, 8 h, and 24 h. We found that ZMD had a high tumor targeting efficiency, with the highest ZMD enrichment at the tumor site at 8 h and partial clearance of ZMD at 24 h (Fig. 5F, H). This may be due to the EPR effect, as well as the release of the drugs from the ZMD in the acidic environment of the tumor, which promotes ROS generation and disrupts the vasculature, which further promotes the accumulation of ZMD. After mice were euthanized, major organs were collected and fluorescence imaging was performed to observe the distribution of ZMD in vivo. The results showed that ZMD was mainly distributed in the liver, kidney and tumor (Fig. 5G, I).

Fig. 5

Efficacy and safety of ZMD in the treatment of tumors in vivo. (A) Schematic illustration of in vivo experimental design. (B) Body weight of mice during different treatments (n = 5, mean ± SD). (C) Tumor images of different groups at the end of the experiment (n = 5). (D) Tumor volumes in different groups at the end of the experiment (n = 5). (E) Tumor weight in different groups at the end of the experiment (n = 5). (F) Quantification of tumor fluorescence intensity of living imaging at different times after tail vein injection (n = 3). (G) Quantification of fluorescence intensity in major organs of mice (n = 3). (H) Living imaging images of tumor-bearing mice after ZMD injection (n= 3). (I) Fluorescence imaging images of major organs and tumor tissues of mice. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

ZMD promotes immune responses in vivo

In our in vitro experiments, we found that ZMD significantly activated the cGAS-STING pathway in HCC and DC cells, promoted DC cell maturation and cytokine expression, and facilitated anti-tumor immunotherapy. Therefore, we performed in vivo experiments to further investigate the effect of ZMD ANPs on anti-tumor immune responses in vivo. Flow cytometry analysis showed that ZMD significantly promoted the maturation of DC cells in mice (Fig. 6A,D). After ZMD treatment, the number of CD8 + T cells and CD4 + T cells in the tumor tissues increased, and macrophages were polarized toward the M1 anti-tumor phenotype (Fig. 6B,C, E–G). Immunofluorescence experiments of tumor tissues further validated that ZMD promoted the infiltration of CD8 + and CD4 + T cells in the tumor environment and reduced macrophage polarization toward the M2 tumor-promoting phenotype (Fig. 6H–K). ZMD promoted the most significant anti-tumor immune response compared to ZIF-8 and ZIF-8@MnCO. Combination of ZMD with radiotherapy significantly activated anti-tumor immunotherapy.

To verify the activation of the cGAS-STING pathway by ZMD in vivo, we performed immunofluorescence experiments of p-TBK1 and p-STING on mouse tumor tissues. The results showed that ZMD significantly promoted the phosphorylation of TBK1 and STING. Compared with ZIF-8 and ZIF-8@MnCO, the activation of the cGAS-STING pathway by ZMD was more pronounced. In addition, the combination of ZMD and radiotherapy significantly activated the cGAS-STING pathway and promoted anti-tumor immune response (Fig. 7A–C). To investigate the effect of ZMD in treating distant metastatic tumors, we constructed a mouse model of tumor lung metastasis. The results showed that ZMD could significantly reduce the number of lung metastatic nodules. The combination of ZMD and radiotherapy could exert a better therapeutic effect on distant metastatic tumors (Fig. 7D,E).

Fig. 6

ZMD NPs promote anti-tumor immune responses in vivo. Flow cytometric representation (A) and quantification (D) of DC cell mutation in different groups of tumor tissues (n = 3). Flow cytometric representation (B) and quantification of CD8 + T cells (E) and CD4 + T cells (F) in different groups of tumor tissues (n = 3). Flow cytometric representation (C) and quantification (G) of macrophages in different groups of tumor tissues. (H) Representative immunofluorescence images of T cells and macrophages in tumor tissue. Scale bar = 50 μm. (I-K) Quantitative analysis of fluorescence intensity in immunofluorescence of CD4 (I), CD8 (J) in T cells, and CD206 in macrophages (K). n = 3. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

Fig. 7

Activation of cGAS-STING pathway and treatment of metastatic lung tumors by ZMD. (A) Representative immunofluorescence images. n = 3. Scale bar = 50 μm. (B-C) Quantitative analysis of p-STING expression (B) and p-TBK1 expression (C) in tumor tissues. n = 3. (D-E) Quantitative analysis (D) and H&E images (E) of different groups of lung metastatic nodules. n = 3. Scale bar = 20 μm. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

Conclusion

In summary, we have engineered and synthesized ZMD, a multifunctional cGAS-STING pathway activator, which can deliver Mn²⁺-Zn²⁺ bipyridyl ions and DOX for activating anti-tumor immunotherapy and augment tumor radiosensitivity. Compared to commercially available cGAS-STING pathway activators, ZMD protects the drug from degradation and specifically targets tumor tissue to release the drug with high biocompatibility. In the tumor environment, ZMD rapidly releases Zn²⁺, Mn²⁺, CO and DOX. The Zn²⁺ and Mn²⁺ irons undergo a Fenton-like reaction with H₂O₂ in the presence of glutathione (GSH), generating ROS that induce oxidative stress and apoptosis in cancer cells. Furthermore, the ROS production enhance tumor cell radiosensitivity. DOX further induces tumor cell death, and the dsDNAs released from cellular damage, synergistically with metal ions, robustly activate the cGAS-STING pathway, inducing anti-tumor immune responses.

Our findings indicate that ZMD NPs significantly augment the radiosensitivity of HCC cells and activate the cGAS-STING pathway, thereby catalyzing a potent anti-tumor immune response. In vitro assays reveal that ZMD synergizes with radiotherapy to suppress tumor cell proliferation, migration, and invasion, while also inducing ROS generation, DNA damage, and apoptosis in tumor cells. In vivo experiments show that ZMD NPs combined with radiotherapy induce substantial tumor regression and resuced metastasis. The activation of the cGAS-STING pathway, both in vitro and in vivo, as evidenced by elevated phosphorylation of STING, TBK1, and IRF3, correlates with DCs maturation and the infiltration of CD8⁺ and CD4⁺ T lymphocytes. These results are consistent with recent studies that highlight the role of oxidative stress and immune activation in improving radiotherapy outcome [35,36,37]. The efficacy of ZMD as a radiosensitizer aligns with findings from similar studies using MOFs and metal-based nanomaterials. For instance, Wu et al. demonstrated that MOF-based delivery systems effectively enhanced drug targeting and reduced systemic toxicity [38]. Additionally, Li et al. reported that metal irons-based NPs combined with immunotherapy enhance tumor cell radiotherapy and significantly inhibit tumor cell growth and metastasis [39]. Iranpour et al. reported that combining metal ions with chemotherapy induces a potent oxidative response, enhancing tumor cell radiosensitivity [39, 40].

Compared to similar studies, we innovatively utilized MOF materials to deliver Mn²⁺-Zn²⁺ bipyridyl ions and DOX in combination with radiotherapy, which significantly activate the cGAS-STING pathway to elicit a potent anti-tumor immune response and induce radiosensitivity in tumor cells [20, 24, 41]. Our study not only underscores the potential of ZMD in enhancing radiotherapy efficacy but also highlights the importance of immune activation in cancer treatment. The dual role of ZMD in inducing ROS-mediated tumor cell death and activating the cGAS-STING pathway offers a comprehensive strategy for HCC treatment. Future research needs to explore the optimization of ZMD synthesis to enhance the performance and safety of the material, develop cost-effective manufacturing processes, and to investigate the potential for clinical translation [42].