PROTAC-based synthetic lethality strategy endogenously activates systemic STING to boost antitumor immunity

Abstract

Activation of the stimulator of interferon genes (STING) pathway drives natural killer (NK) cells and T cells to orchestrate multidimensional antitumor immune responses. While cytosolic DNA accumulation represents a superior endogenous strategy for STING activation, DNA repair machinery substantially constrains its immunogenic potential. Here, we propose a promising therapeutic strategy that leverages proteolysis-targeting chimera (PROTAC)–mediated degradation of PARP1 [poly(ADP-ribose) polymerase 1] and BRD4 (bromodomain-containing protein 4) to induce synthetic lethality, thereby disrupting DNA repair machinery that drives nuclear-to-cytosolic DNA leakage, surpassing the STING activation threshold to ignite cGAS-STING–mediated innate immunity. Our strategy demonstrates superior antitumor efficacy across multiple tumor models, eliciting robust CD8⁺ T cell– and NK cell–mediated immunity while suppressing pulmonary metastasis progression. This strategic integration of synthetic lethality with an immunogenic stress response establishes a previously unidentified paradigm for expanding broad applications by cGAS-STING–mediated innate immunity.

PROTAC co-degrades PARP1/BRD4, disrupting the DNA repair process, activating the STING pathway via cytosolic DNA accumulation.

INTRODUCTION

Stimulator of interferon genes (STING) is a pivotal innate immune receptor that mediates cytoplasmic DNA sensing through intracellular signaling cascades (1–3). Upon activation, STING promotes the robust secretion of type I interferons such as interferon-β (IFN-β) (4, 5), along with a spectrum of pro-inflammatory cytokines (6), which drives multifaceted antitumor immune functions by recruiting and activating immune cells and enhancing natural killer (NK) cell– and T cell–mediated tumor killing (7–9). Although the STING pathway has emerged as a pivotal target in cancer immunotherapy, current exogenous activation strategies based on this pathway still exhibit notable limitations. As commonly used STING agonists, cyclic dinucleotides are currently restricted to intratumoral administration owing to their poor bioavailability and stability (6, 10–14). To address the limitations of intratumoral delivery, researchers have engineered nonnucleotidic small-molecule STING agonists, exemplified by SR-717 and MSA-2, which demonstrate that systemic administration of these compounds effectively induces antitumor immune responses (15–18). However, the rational design of new small-molecule STING agonists remains a formidable challenge because of structural complexity in balancing STING isoform activation, systemic exposure, and tumor-selective immunomodulation. Therefore, evolving endogenous activation strategies has become imperative to transcend current therapeutic plateaus.

Cytosolic DNA serves as the endogenous native activator of the STING pathway, dictating the magnitude and specificity of intracellular immune responses (19–21). Strategic accumulation of cytoplasmic DNA fragments may provoke STING pathway activation through controlled genomic destabilization, representing an intervention strategy that bypasses current pharmacological limitations. The cytosolic accumulation of DNA primarily originates from nuclear DNA damage–derived fragments (22, 23); however, the DNA damage response system dynamically orchestrates DNA repair processes through homeostatic surveillance mechanisms, which tightly restrict cytosolic DNA accumulation below the activation threshold of the STING pathway (24, 25). Hence, targeted disruption of DNA repair machinery can induce nuclear-to-cytosolic translocation of damaged DNA fragments, thereby overcoming the physiological activation threshold of the STING pathway. Poly(ADP-ribose) polymerase 1 (PARP1), as a protein involved in DNA damage repair and the maintenance of genomic integrity, exhibits synthetic lethality through its inhibitors (PARP inhibitors) in homologous recombination (HR) deficiency–positive tumors harboring BRCA1/2 mutations. Mechanistically, PARP inhibitors stabilize PARP-DNA complexes at single-strand breaks, preventing their resolution and leading to replication fork collapse that generates lethal double-strand breaks (26–29). The resultant cytosolic accumulation of unrepaired DNA fragments potently activates STING-mediated innate immune signaling and subsequent antitumor immunity (30–32). However, the clinical utility of this strategy is constrained by the fact that only ~15% of malignancies exhibit intrinsic HR deficiency. To address this limitation, recent investigations have focused on bromodomain-containing protein 4 (BRD4), an epigenetic regulator governing DNA repair gene networks. Pharmacological inhibition of BRD4 down-regulates critical HR components, thereby inducing a therapeutically exploitable “acquired HR-deficient (HRD)” phenotype (33). Cotargeting degradation of PARP1 and BRD4 thus represents a rational strategy to induce synthetic lethality via dual blockade of the DNA repair pathway, with resultant cytosolic DNA leakage activating the STING signaling cascade to execute antitumor effects.

Current strategies for intracellular protein modulation primarily rely on small-molecule inhibitors and emerging proteolysis-targeting chimeras (PROTACs). While conventional inhibitors suffer from transient target engagement, heterogeneous therapeutic responses, and adaptive resistance resulting from their occupancy-driven pharmacology, PROTACs leverage catalytic ternary complex formation (target-PROTAC-E3 ligase) to hijack the ubiquitin-proteasome system, enabling sustained protein degradation through a dynamic, event-driven mechanism that bypasses traditional pharmacodynamic constraints (34–37). Herein, we propose a promising therapeutic strategy that simultaneously degrades PARP1 and BRD4 to achieve synthetic lethality, thereby disrupting DNA repair machinery to promote cytosolic accumulation of fragmented double-stranded DNA (dsDNA), which activates the STING pathway in situ. We developed two distinct PROTACs through structure-guided rational design, specifically engineered to selectively degrade PARP1 and BRD4. This combinatorial degradation of PARP1 and BRD4 triggers dual repair system collapse in tumor cells: PARP1 ablation disrupts the base excision repair pathway, converting unrepaired single-strand breaks into replication fork–derived lethal double-strand breaks during DNA replication, while BRD4 degradation epigenetically silences transcription of HR core components, abolishing compensatory DNA repair capacity. The resultant genomic instability drives substantial DNA leakage into cytosol through either nuclear envelope breach or vesicular transport dysregulation. Cytosolic DNA fragments are recognized by cyclic GMP (guanosine 5′-monophosphate)-AMP (adenosine 5′-monophosphate) synthase (cGAS) as danger signals, initiating cGAS-STING signaling pathway activation that culminates in type I interferon (IFN-β) production and antigen-presenting cell (APC) activation (Fig. 1). Overall, this study establishes a synergistic framework wherein synthetic lethality is coupled with immunogenic stress responses to drive tumor cell eradication. Our findings highlight the therapeutic potential of exploiting endogenous STING activation via rational PROTAC design, offering a versatile platform for next-generation cancer immunotherapy.

Fig. 1. Schematic illustration for PRO_P+B-induced cytoplasmic DNA accumulation to activate the cGAS-STING signaling pathway for immunotherapy.

(A) Structural formulas and mechanisms of action of PRO_P and PRO_B. (B) PRO_P+B induces synthetic lethality in tumor cells by mediating the degradation of PARP1 and BRD4, thereby disrupting DNA repair mechanisms. This process leads to the leakage of nuclear DNA into the cytoplasm, subsequently activating the cGAS-STING–mediated innate immune response.

RESULTS

PRO_P+B potently degrades proteins and induces DNA damage

We synthesized two PROTACs by conjugating the BRD4 inhibitor JQ1 and PARP1 inhibitor olaparib with E3 ligase–recruiting moieties, designated as PRO_B and PRO_P, respectively (Fig. 2A). The successful synthesis of PRO_B and PRO_P was verified by ¹H nuclear magnetic resonance spectroscopy and electrospray ionization mass spectrometry analyses, respectively (figs. S1 to S7). Initial Western blotting (WB) analysis revealed potent degradation of both targets in B16F10 cells, achieving half-maximal degradation concentration values of 0.12 μM for each PROTAC (fig. S8). The combinatorial treatment with PRO_P+B induced concentration-dependent dual protein degradation of PARP1 and BRD4, achieving >80 and >50% reduction at 0.01 μM, respectively, and reaching a maximal degradation efficiency of 90% for both targets at 10 μM (Fig. 2, B and C). This complete target eradication at 10 μM prompted its selection for subsequent assays. Immunofluorescence quantification confirmed these findings through marked reduction of BRD4-associated fluorescence and PARP1-specific signals in B16F10 cells (Fig. 2, D and E, and fig. S9). Crucially, the universal efficacy of PRO_P+B extended across multiple murine cell lines, including 4T1 (breast), H22 (hepatoma), KPC (pancreatic), and CT26 (colorectal) cell lines, where near-complete fluorescence signal loss confirmed robust, pancancer target engagement (Fig. 2, F to H). These orthogonal validation approaches conclusively establish PRO_P+B as a robust dual-degrader platform with broad tumor cell applicability.

Fig. 2. PRO_P+B induced dual PARP1/BRD4 depletion, exacerbating DNA damage.

(A) Schematic representation of PROTAC design targeting PARP1 and BRD4. (B) WB analysis of BRD4 and PARP1 protein levels in B16F10 cells treated with PRO_P, PRO_B, and PRO_P+B for 12 hours. (C) WB analysis of BRD4 and PARP1 protein levels in B16F10 cells treated with PRO_P+B for 24 hours at indicated concentrations. Quantification of BRD4 (D) and PARP1 (E) expression levels in B16F10 cells after incubating with PBS, PRO_P, PRO_B, and PRO_P+B for 24 hours. a.u., arbitrary units. BRD4 (F) and PARP1 (G) expression detected by immunofluorescence in 4T1, H22, KPC, and CT26 cells after incubating with PRO_P+B for 24 hours. The signals of BRD4 and PARP1 were generated by Alexa Fluor 488– and Alexa Fluor 647–conjugated secondary antibodies, respectively. (H) Heatmap visualization of target protein expression levels after incubating with PRO_P+B for 24 hours. (I) Schematic diagram of PROTAC preventing DNA damage repair. (J) WB analysis of γ-H2AX protein levels in B16F10 cells treated with PRO_P, PRO_B, and PRO_P+B for 24 hours. (K) Comet assay for B16F10 cells under diverse treatments. (L) Immunofluorescence of γ-H2AX staining for DNA fragmentation visualization after incubation of PRO_P+B for 24 hours in 4T1, H22, KPC, and CT26 cells. The signals of γ-H2AX were generated by Alexa Fluor 488–conjugated secondary antibodies. (M) Comet assay for 4T1, H22, KPC, and CT26 cells after PRO_P+B treatments. (N) DNA damage was expressed by the ratio of comet tail on the basis of the comet assay in (M). Results are presented as the means ± SD.

BRD4, a master regulator of superenhancer architecture and oncogenic transcription programs (38), drives tumorigenesis through the coordinated modulation of DNA replication and damage response pathways. Targeted degradation of BRD4 disrupts HR repair by down-regulating key effectors such as BRCA1 and RAD51 expression—critical mediators of DNA damage tolerance (33, 39). This pharmacological induction of HR deficiency creates therapeutic vulnerability through persistent replication stress, ultimately manifesting as genomic instability and chromosomal aberrations. Our PROTAC-based strategy (PRO_P+B cotreatment) amplifies this synthetic lethal effect by concurrently targeting PARP1-mediated DNA repair pathways (Fig. 2I). To quantify DNA damage dynamics, the γ-H2AX formation was monitored, which is the gold-standard biomarker for double-strand break detection. Integrated analysis through immunofluorescence staining and WB quantification demonstrated that 10 μM PRO_P+B combination therapy significantly increased γ-H2AX expression in B16F10 cells compared to single-agent treatments (Fig. 2J and fig. S10). Time-course analysis demonstrated progressive γ-H2AX accumulation, reaching maximal signal intensity at 24 hours posttreatment (fig. S11). Complementing these findings, single-cell comet assays showed notable differences in nuclear morphology: While phosphate-buffered saline (PBS)– or monotherapy-treated cells maintained an intact chromatin structure (tail moment <5%), PRO_P+B exposure generated pronounced comet tails, indicative of severe DNA fragmentation (Fig. 2, K to N). Notably, the pancancer efficacy of this DNA damage amplification was consistently validated across four distinct cell lines (4T1, H22, KPC, and CT26). Results revealed that PRO_P+B treatment universally increased γ-H2AX–positive nuclei by >3.0-fold compared to controls, with comet tailing percentages exceeding 20% in all tested lines (Fig. 2, M and N, and fig. S12). These results further underscore the potency of the PRO_P and PRO_B combination in inducing persistent, widespread DNA damage across various tumor contexts, thereby validating this dual-targeting approach as a broadly effective anticancer strategy.

Cytosolic DNA release

Given the profound DNA damaging ability induced by PRO_P+B, we next examined the downstream accumulation of cytosolic dsDNA, a pivotal trigger of the innate immune system. To characterize PRO_P+B-induced cytoplasmic dsDNA accumulation, we performed immunofluorescence staining using an anti-dsDNA antibody. Tumor cells treated with PRO_P, PRO_B, or PRO_P+B were analyzed accordingly. Consistent with our hypothesis, PRO_P+B treatment induced significant cytoplasmic dsDNA aggregation in B16F10 cells (Fig. 3A). In contrast, monotherapy groups (PRO_P or PRO_B alone) suppressed dsDNA fragment generation. This dsDNA accumulation phenotype was recapitulated in four additional cell lines (fig. S13). Quantitative analysis revealed that while monotherapy with either PRO_P or PRO_B caused marginal increases (3.7- and 2.9-fold versus PBS, respectively), the PROTAC combination triggered massive cytosolic dsDNA accumulation in B16F10 cells, reaching 12.6-fold elevation over controls (P < 0.0001; Fig. 3B). This dsDNA efflux exhibited pancancer significance, with 4T1 (9.0-fold), H22 (8.9-fold), KPC (8.6-fold), and CT26 (9.5-fold) cells all showing significant cytosolic dsDNA enrichment post–PRO_P+B treatment (Fig. 3C).

Fig. 3. Damaged cytosolic dsDNA activates the STING signaling pathway.

(A) dsDNA expression levels detected by immunofluorescence in B16F10 cells after incubating with indicated treatments (PBS, PRO_P, PRO_B, and PRO_P+B) for 24 hours. The signals of dsDNA were generated by Alexa Fluor 488–conjugated secondary antibodies. Quantification of dsDNA in the cytoplasm of the B16F10 cells (B) and 4T1, H22, KPC, and CT26 cells (C). n = 3 biologically independent samples. (D) Schematic diagram of STING pathway activation upon recognition of cytosolic dsDNA. ELISA revealed cGAMP production in B16F10 cells (E) and 4T1, H22, KPC, and CT26 cells (G) 24 hours after indicated treatments (PBS, PRO_P, PRO_B, and PRO_P+B). (F) ELISA revealed cGAMP production in B16F10 cells after incubation of PRO_P+B with or without DNase I addition. (H) WB analysis of cGAS, p-STING, and p-IRF3 protein levels in B16F10 cells treated with PBS, PRO_P, PRO_B, and PRO_P+B for 24 hours. ELISA revealed IFN-β production in B16F10 cells (I) and 4T1, H22, KPC, and CT26 cells (J) 24 hours after indicated treatments (PBS, PRO_P, PRO_B, and PRO_P+B). (K) PD-L1 expression levels detected by immunofluorescence in B16F10 cells after incubating with indicated treatments (PBS, PRO_P, PRO_B, and PRO_P+B) for 24 hours. The signals of PD-L1 were generated by Alexa Fluor 488–conjugated secondary antibodies. (L) Flow cytometry analysis of PD-L1 expression in 4T1, H22, KPC, and CT26 cells treated with indicated treatments (PBS, PRO_P, PRO_B, and PRO_P+B) for 24 hours. (K) is the quantification of (M). Results are presented as the means ± SD.

cGAS-STING pathway stimulation in tumor cells

Building upon the observed cytosolic dsDNA accumulation, we systematically mapped the downstream immune activation cascade through a series of experiments (Fig. 3D). 2′3′-Cyclic GMP-AMP (cGAMP) is an intracellular second messenger that is synthesized in response to cytosolic dsDNA and activates the innate immune STING pathway (40). Quantitative 2′,3′-cGAMP enzyme-linked immunosorbent assay (ELISA) revealed that PRO_P+B treatment induced a 3.5-fold increase in cyclic dinucleotide levels compared to PBS controls in B16F10 cells, with statistical significance confirmed through rigorous analysis (Fig. 3E). This response demonstrated both dose dependency and temporal dynamics, reaching maximal cGAMP accumulation at 60 hours posttreatment (fig. S14). Notably, coadministration with deoxyribonuclease (DNase) I attenuated cGAMP production by 82.5%, establishing dsDNA as the principal ligand driving cGAS activation (Fig. 3F). A pancancer analysis demonstrated robust cGAMP elevation across multiple models, including 4T1 (2.8-fold), H22 (2.0-fold), KPC (3.9-fold), and CT26 (3.4-fold), confirming the broad applicability of this pathway engagement (Fig. 3G). Integrated WB analysis across five tumor cell models (B16F10, 4T1, H22, KPC, and CT26) revealed that PRO_P+B combination therapy markedly up-regulated cGAS protein expression, with concurrent accumulation of phosphorylated STING (p-STING) and phosphorylated interferon regulatory factor 3 (p-IRF3) (Fig. 3H and fig. S15). This sequential phosphorylation cascade, characterized by enhanced cGAS expression followed by STING and IRF3 phosphorylation, precisely aligns with canonical STING pathway activation mechanisms. Subsequent ELISA quantification confirmed that IFN-β secretion increased 2.0- to 3.1-fold across cell lines (Fig. 3, I and J, and fig. S16). Collectively, this integrated dataset establishes a complete sequence of events: PRO_P+B-induced genomic instability triggers cytosolic dsDNA release, which promotes cGAS oligomerization and subsequent cGAMP synthesis. This second messenger then activates cGAS-STING-IRF3 phosphorylation cascades, ultimately driving type I interferon production. These findings establish a dual role for PROTAC-mediated DNA damage as both a cytotoxic event and immunogenic stimulus, thereby creating a functional bridge between targeted protein degradation and innate immune activation.

Substantial intratumoral accumulation of cGAMP has been reported to up-regulate programmed death-ligand 1 (PD-L1) in tumor cells, impeding the initiation and activation of the antitumor immune response (41, 42). This paradoxical response aligns with recent findings that PARP1 inhibition stabilizes PD-L1 via glycogen synthase kinase 3β inactivation, posing therapeutic limitations (43). Notably, the small-molecule BRD4 inhibitor JQ1 has been shown to attenuate immune evasion by down-regulating PD-L1 expression (44–46). We therefore quantified PD-L1 expression dynamics across PROTAC regimens using multimodal validation. In B16F10 cells, PARP1 degradation via PRO_P monotherapy increased PD-L1 surface expression by 1.7-fold versus PBS-treated controls (fig. S17). Selective BRD4 degradation through PRO_B or PRO_P+B treatment counteracted this effect, reducing PD-L1 levels to 49.5 and 35.5% of PBS controls, respectively. Confocal microscopy with anti–PD-L1-Alexa Fluor 647 confirmed these findings, showing membrane-localized PD-L1 reduction in >58% of PRO_P+B-treated cells compared with PRO_P (Fig. 3K and fig. S18). This trend was consistently observed across multiple tumor lineages, with 4T1, H22, KPC, and CT26 cells showing similar PD-L1 elevation after PRO_P treatment compared to PBS controls (Fig. 3, L and M). Notably, combinatorial treatment with PRO_P+B completely reversed this immune checkpoint potentiation, achieving statistically significant suppression of PD-L1 expression. Therapeutic synergy emerged from combining these orthogonal effects: While PARP1 degradation amplified cGAMP-driven STING activation, concurrent BRD4 ablation suppressed PD-L1 through bromodomain-dependent transcriptional regulation. Given that the PD-1/PD-L1 axis is a negative regulatory signaling pathway that can trigger multiple immunosuppressive effects, including T cell anergy, the down-regulation of PD-L1 mediated by PRO_P+B is expected to enhance T cell function and proliferation.

cGAS-STING pathway stimulation in immune cells

As professional APCs, dendritic cells (DCs) initiate the adaptive immune response by ingesting, processing, and presenting antigens to naive T cells, thereby activating T cells (47, 48). Effective antigen presentation by DCs requires their maturation, a process characterized by the up-regulation of costimulatory molecules, including type I interferons such as IFN-β, and the secretion of pro-inflammatory cytokines (49). Previous experimental results have shown that after stimulation with PRO_P+B, the STING pathway in tumor cells is activated, leading to the secretion of a large amount of IFN-β. On the basis of this, we hypothesized that tumor cells treated with PRO_P+B, because of the presence of a large amount of IFN-β and dsDNA, may induce the activation of the STING pathway in immune cells and the maturation of DCs when co-incubated with DCs (Fig. 4A). To test this hypothesis, we used bone marrow–derived DCs (BMDCs) isolated from C57BL/6J mice to analyze the promoting effect of PRO_P+B on BMDC maturation in vitro. Specifically, B16F10 cells were pretreated with PRO_P, PRO_B, or PRO_P+B and then co-incubated with BMDCs. Flow cytometry was performed to count mature BMDCs (CD11c⁺CD80⁺CD86⁺, denoted as mDC) (Fig. 4B and fig. S19). The results showed that there was no significant difference in the number of mDCs in BMDCs incubated with B16F10 cells treated with PRO_B compared with the control group (BMDCs incubated with untreated cells). However, the number of mDCs increased to ~41.9% in BMDCs incubated with cells treated with PRO_P and to 57.1% in BMDCs incubated with cells treated with PRO_P+B. To further validate our findings, we isolated bone marrow–derived macrophages (BMDMs) and repeated the aforementioned experiments. The results demonstrated that tumor cells treated with PRO_P+B significantly up-regulated the surface expression of CD86 on BMDMs, indicating enhanced APC maturation (fig. S20). These findings collectively confirm that PRO_P+B-stimulated tumor cells have a broad capacity to promote APC activation, reinforcing the immunomodulatory potential of this therapeutic approach.

Fig. 4. STING activation and DC maturation by tumor-derived cytoplasmic DNA.

(A) Schematic diagram of STING pathway activation and maturation of DCs. (B) Representative flow cytometric analysis image and relative quantifications of mature DC markers CD80/CD86 in BMDCs co-incubated with B16F10 cells treated with PBS, PRO_P, PRO_B, and PRO_P+B for 24 hours. (C) Schematic diagram of STING pathway activation and maturation of DCs. Production of cGAMP in DC2.4 cells (D) and THP1 cells (E), which were co-incubated with B16F10, 4T1, H22, KPC, and CT26 cells after the tumor cells were treated with PBS, PRO_P, PRO_B, and PRO_P+B, respectively, measured by ELISA assay. (F) WB analysis of cGAS, p-STING, and p-IRF3 protein levels in DC2.4 cells that were co-incubated with B16F10, 4T1, H22, KPC, and CT26 cells after the tumor cells were treated with PBS, PRO_P, PRO_B, and PRO_P+B, respectively. Results are presented as the means ± SD.

To evaluate the potential of PRO_P+B-induced immunomodulation, we interrogated STING pathway activation in both murine DCs (DC2.4) and human monocytic cells (THP-1) using a transwell coculture system (Fig. 4C). Tumor cells pretreated with PRO_P+B were physically segregated from immune cells to isolate paracrine signaling effects. Quantitative 2′,3′-cGAMP ELISA demonstrated that PRO_P+B-treated B16F10 cells induced substantial 4.6- and 11.2-fold increases in this cyclic dinucleotide in DC2.4 and THP-1 cells, respectively, compared to PBS controls (Fig. 4, D and E). This response exhibited similar trends across all five cancer cell lines. WB analysis of DC2.4 cells exposed to PRO_P+B-treated tumor lysates revealed coordinated up-regulation of the key proteins (cGAS, p-STING, and p-IRF3) in the STING signaling pathway relative to untreated controls (Fig. 4F). This phosphorylation cascade confirms complete pathway engagement from cytosolic DNA sensing through interferon regulatory factor activation.

The mechanistic convergence observed between murine and human cell lines underscores PRO_P+B as a robust activator of cross-species STING signaling. Tumor cells subjected to PRO_P+B treatment generate two critical immunogenic outputs: cytosolic dsDNA fragments that serve as cGAS ligands and paracrine cGAMP capable of diffusing into neighboring APCs. This dual-action mechanism not only amplifies innate immune sensing but also integrates with previously characterized effects, including PD-L1 down-regulation and DC maturation, thereby establishing PRO_P+B as a multifunctional modulator of antitumor immunity.

PRO_P+B showed synthetic lethality in multiple cancer lineages

To ensure synchronized pharmacokinetics and enhance tumor-specific accumulation of both PROTAC molecules during systemic circulation, a carrier system was indispensable. Among various nanoplatforms, lipid nanoparticles (liposomes) represent a well-established and clinically validated drug delivery system with demonstrated advantages in improving drug stability, bioavailability, and targeted delivery. Capitalizing on these clinically proven benefits, we developed a dual PROTAC-encapsulated lipid nanoparticle platform (LNP@PRO_P+B) using an optimized solvent evaporation-hydration method. The formulation integrates DOPC (1,2-dioleoyl-sn-glycero-3-phosphocholine) as the primary phospholipid (60 mol %), cholesterol (35 mol %) for membrane stabilization, and DSPE-PEG₂₀₀₀ (1,2-distearoyl-sn-glycero-3-phosphoethanolamine-polyethylene glycol; 5 mol %) to enhance systemic circulation. High-performance liquid chromatography quantification demonstrated encapsulation efficiencies of 39.6 ± 2.1% for PRO_P and 25.3 ± 1.8% for PRO_B, with corresponding drug loading capacities reaching 13.2 ± 0.7 and 6.72 ± 0.4% (w/w). Dynamic light scattering analysis revealed monodisperse nanoparticles with a hydrodynamic diameter of 120 ± 10 nm (polydispersity index, 0.19 ± 0.02; parameters optimized for enhanced permeability and retention–mediated tumor targeting; fig. S21). The formulation exhibited excellent colloidal stability in physiological buffer (PBS, pH 7.4) over 7 days, maintaining the particle size within 15% of initial measurements and polydispersity index below 0.25.

Building upon the foundational synthetic lethality paradigm of PARP1 inhibition in HRD tumors, we hypothesized that BRD4 degradation could induced an HRD phenotype, thereby sensitizing cancer cells to PRO_P-mediated PARP1 trapping. This hypothesis was systematically evaluated through complementary cytotoxicity assays spanning nine human and murine cancer models. Quantitative MTT [3-(4,5-dimethyl thiazol-2-yl)-2,5-diphenyltetrazolium bromide] analysis revealed strong synergistic interactions between LNP@PRO_P and LNP@PRO_B, with combination index (CI) values <1 across all tested cell lines (Fig. 5A and fig. S22). Notably, the therapeutic synergy was particularly pronounced in BRCA1/2-wild-type models, suggesting that BRD4 degradation expands PARP1 inhibitor (PARP1i) sensitivity beyond canonical HRD contexts. Functional validation using clonogenic survival assays confirmed this synthetic lethality. Compared to PBS, LNP@PRO_P+B reduced colony-forming units by 95.5 ± 4.0% in B16F10 cells versus 42.5 ± 8.7% (LNP@PRO_B monotherapy) and 31.7 ± 10.5% (LNP@PRO_P monotherapy) (Fig. 5B). A similar trend was observed across four additional tumor types (4T1: 55.2 ± 3.7% reduction; H22: 95.6 ± 2.3%; KPC: 93.0 ± 2.4%; CT26: 95 ± 4.7%) (Fig. 5, C and D). Collectively, these findings delineate a three-phase synthetic lethality mechanism: BRD4 degradation down-regulates HR repair effectors, PARP1 trapping induces replication fork collapse, and concomitant HR impairment prevents damage resolution. The BRCA-wild-type models suggest that this approach could overcome current limitations of PARP1i therapy.

Fig. 5. Systemic LNP@PRO_P+B treatment exerts a potent antitumor effect.

(A) IC₅₀ (median inhibitory concentration) and CI values of different cell lines after incubation with LNP@PRO_P, LNP@PRO_B, and LNP@PRO_P+B. (B and C) Representative pictures of clonogenic assay in B16F10, 4T1, H22, KPC, and CT26 cells treated with PBS (G1), LNP@PRO_P (G2), LNP@PRO_B (G3), or LNP@PRO_P+B (G4) for 24 hours. (D) Colony-forming efficiency of B16F10, 4T1, H22, KPC, and CT26 cells. (E) Schematic illustration of the timeline for B16F10 tumor inoculation and treatments in a C57BL/6 mouse model. Mice with ~100-mm³ subcutaneous tumors were intravenously (iv) administered with LNP@PRO_P, LNP@PR_B, and LNP@PRO_P+B five times at a PRO_B equivalent dose of 10 mg kg⁻¹. sc, subcutaneously. (F) Tumor growth of B16F10 tumor–bearing mice treated with LNP@PRO_P, LNP@PRO_B, or LNP@PRO_P+B. n = 7 biologically independent samples. (G) Survival of B16F10 tumor–bearing mice treated with LNP@PRO_P, LNP@PRO_B, or LNP@PRO_P+B. (H) Immunofluorescence staining of BRD4, PARP1, PD-L1, and γ-H2AX expression levels in B16F10 tumor after indicated treatment. (I) WB analysis of cGAS, p-STING, and p-IRF3 protein levels in B16F10 tumor after indicated treatment. (J) Schematic illustration of the timeline for 4T1/CT26 tumor inoculation and treatments in a BALB/C mouse model. Tumor growth (K) and survival (L) of 4T1 tumor–bearing mice treated with LNP@PRO_P, LNP@PRO_B, or LNP@PRO_P+B. n = 7 biologically independent samples. Tumor growth (M) and survival (N) of CT26 tumor–bearing mice treated with PBS (G1), LNP@PRO_P (G2), LNP@PRO_B (G3), or LNP@PRO_P+B (G4). n = 7 biologically independent samples. Results are presented as the means ± SEM. (O) Schematic illustration of the timeline for B16F10 tumor inoculation and treatments in a NCG mouse model. (P) Tumor growth of B16F10 tumor–bearing mice treated with PBS or LNP@PRO_P+B. n = 6 biologically independent samples. (Q) Photographs of the excised tumors. (R) Tumor weights of different groups after treatments. (S) Tumor inhibition of different groups after treatments. Results are presented as the means ± SD.

To further validate the necessity of using PROTACs over conventional inhibitors, we established a B16F10-Luc melanoma lung metastasis model in C57BL/6 mice and initiated treatment via tail vein injection of either LNP@PRO_P+B or LNP@PARP1i + BRD4i starting from day 14 post–tumor inoculation, continuing for five consecutive days (fig. S23A). The results demonstrated that LNP@PRO_P+B significantly suppressed both pulmonary metastatic lesions and primary tumor growth, in marked contrast to the PBS and LNP@PARP1i + BRD4i groups, both of which exhibited extensive lung metastasis with strong bioluminescent signals (fig. S23, B and C). This comparative experiment unequivocally confirmed the superior efficacy of PROTACs over an inhibitor. Next, we used three orthogonal tumor models characterized by distinct genetic and immunological profiles to evaluate the therapeutic potential of LNP@PRO_P+B in vivo (Fig. 5E). First, the poorly immunogenic B16F10 melanoma model was used, with treatments administered via tail vein injection starting 7 days after tumor inoculation and repeated for 5 consecutive days. As shown in Fig. 5F, while tumors in the LNP@PRO_P and LNP@PRO_B monotherapy groups exhibited substantial growth, the dual-PROTAC combination (LNP@PRO_P+B) significantly delayed tumor progression, maintaining tumor volumes below 300 mm³ throughout the observation period and demonstrating superior antitumor efficacy. A long-term survival monitoring experiment demonstrated that LNP@PRO_P+B was superior to other control groups in terms of prolonged survival: All control mice succumbed within 30 days, and monotherapy groups (LNP@PRO_P or LNP@PRO_B) survived up to 40 days, while the LNP@PRO_P+B group achieved 42.9% 60-day survival rate (three of seven mice, P < 0.001) (Fig. 5G). Mechanistic dissection at day 7 posttreatment demonstrated target-specific pharmacodynamics. Immunofluorescence staining analysis revealed 45.5 ± 7.1% BRD4 degradation and a 78.9 ± 3.8% PARP1 level decrease in LNP@PRO_P+B-treated tumor (Fig. 5H and fig. S24). Notably, LNP@PRO_P-induced PD-L1 up-regulation (1.3 ± 0.2–fold versus control) was abrogated by co-delivery of PRO_B, suggesting that BRD4 degradation counteracts adaptive immune resistance. γ-H2AX immunofluorescence confirmed the most pronounced DNA damage in the LNP@PRO_P+B group. WB analysis demonstrated that LNP@PRO_P+B potently activated the cGAS-STING pathway, with cGAS, p-STING, and p-IRF3 levels up-regulated by 3.6-, 3.8-, and 3.5-fold compared to PBS, respectively (Fig. 5I).

To systematically evaluate the therapeutic efficacy of LNP@PRO_P+B across distinct immunological contexts, we established tumor models using immunologically divergent 4T1 and CT26 cell lines (Fig. 5J). The CT26 model represents immunologically “hot” tumors, whereas 4T1 tumors exhibit a “cold” phenotype with marked resistance to immune checkpoint blockade. In the 4T1 model, LNP@PRO_P+B treatment achieved 69.8 ± 7.5% tumor growth inhibition and extended median survival from 28 to 38 days compared to controls, and 20% of mice attained long-term survival at the 60-day end point (Fig. 5, K and L, and fig. S25). Notably, the CT26 model demonstrated superior responses, with 30% maintaining tumor-free status at study termination (Fig. 5, M and N). Mechanistically, LNP@PRO_P+B induced robust degradation of PARP1 and BRD4 in both models, concurrently reducing PD-L1 expression to 83.7 ± 7.1% in 4T1 and 30.2 ± 12.9% in CT26 tumors (figs. S26 and S27). Substantial DNA damage was evidenced by 1.4-fold (4T1) and 3.2-fold (CT26) up-regulation of γ-H2AX. Posttreatment analysis revealed significant activation of the STING pathway, characterized by enhanced cGAS, p-STING, and p-IRF3 secretion, suggesting that sustained innate immune activation contributes to therapeutic efficacy (fig. S28). Together, these findings not only elucidate the dual mechanisms of target protein degradation and STING pathway activation underlying therapeutic effects of LNP@PRO_P+B but also highlight the critical influence of tumor immunogenicity on treatment outcomes. Furthermore, longitudinal monitoring of murine body weights throughout the therapeutic regimen revealed no statistically significant fluctuations at the implemented dosage (PRO_P+B: 12 mg kg⁻¹, five cycles), supporting the favorable tolerability of this combinatorial nanotherapeutic strategy (fig. S29).

To achieve sustained tumor eradication and prevent recurrence, the deep engagement of the host immune system is typically required to establish long-lasting antitumor responses. To rigorously evaluate the critical role of the immune system, we established tumor models in severely immunodeficient NCG mice, which lack functional T cells, B cells, and NK cells (Fig. 5O). These mice were treated with the same regimen (LNP@PRO_P+B) that demonstrated significant efficacy in immunocompetent C57BL/6 mice. As shown in Fig. 5 (P to R), tumor growth was initially suppressed comparably to that in immunocompetent mice (tumor volume remained below 300 mm³) within the first week of treatment. However, tumors in NCG mice resumed progressive growth in the second week, ultimately exhibiting no significant difference in volume compared to the PBS control group by the end of the study. The tumor inhibition rate in NCG mice was only 33.1 ± 7.8%, significantly lower than the 88.1 ± 7.4% observed in immunocompetent C57BL/6 mice (Fig. 5S). This marked contrast confirms that functional immune cells, particularly those involved in adaptive immunity, are indispensable for the durable therapeutic effects and sustained tumor control achieved through our synthetic lethality strategy combined with STING agonist treatment.

Systemic STING pathway activation for antitumor immunotherapy

Given the previously demonstrated activation of the cGAS-STING pathway in vivo, we hypothesized that this pathway activation might facilitate DC maturation within tumor-draining lymph nodes, thereby inducing subsequent T cell activation (Fig. 6A). To investigate whether LNP@PRO_P+B treatment could induce DC maturation in vivo, we isolated lymph nodes and prepared single-cell suspensions. Cells were stained with fluorescein isothiocyanate (FITC)–conjugated anti-CD11c and phycoerythrin (PE)–conjugated anti-CD86 antibodies for flow cytometric analysis of CD11c⁺CD86⁺ DC populations (gating strategy shown in fig. S30). Notably, the LNP@PRO_P+B treatment group exhibited the highest proportion of CD11c⁺CD86⁺ DCs (18.7%) in tumor-draining lymph nodes of B16F10 tumor–bearing mice, representing a 3.0-fold increase compared to the PBS control group and significantly exceeding the level observed in both LNP@PRO_P and LNP@PRO_B groups (Fig. 6B). These results suggest that LNP@PRO_P+B robustly promotes DC maturation, likely through the synergistic enhancement of tumor-derived dsDNA–induced cGAS-STING pathway activation.

Fig. 6. Antitumor immune response activation by LNP@PRO_P+B.

(A) Schematic diagram of immune system activation. (B) Representative flow cytometric analysis and relative quantifications of mature DCs in tumor-draining lymph node cells after indicated treatment. (C) Analysis of B16F10 tumors after 21 days of treatment for effector cytokine production by intratumoral CD8⁺ T cells. Representative flow cytometric analysis images (D) and relative quantifications (E) of IFN-γ⁺ CD8⁺ and GzmB⁺ CD8⁺ T cells in the tumor. TILs, tumor-infiltrating lymphocytes. (F and G) Analysis of spleen in B16F10 tumor–bearing mice after 21 days of treatment for effector cytokine production by CD8⁺ T cells. (H and I) Analysis of spleen in B16F10 tumor–bearing mice after 21 days of treatment for effector cytokine production by NK⁺ cells. (J) Analysis of B16F10 tumors after 21 days of treatment for effector cytokine production by intratumoral NK⁺ cells. Representative flow cytometric analysis images (K) and relative quantifications (L) of IFN-γ⁺ NK⁺ and GzmB⁺ NK⁺ cells in the tumor. (M to O) Representative flow cytometric analysis and quantification of CD4⁺ CD25⁺ Foxp3⁺ cells in the tumor tissues and spleen. Results are presented as the means ± SD.

The augmented maturation of DCs creates favorable conditions for efficient antigen presentation during subsequent phases of the immune response. We therefore further investigated the proportion of cytotoxic T lymphocytes (CD3⁺CD8⁺ T cells) in both tumor tissues and spleens posttreatment (gating strategy shown in fig. S31). Notably, treatment with LNP@PRO_P+B generated the highest tumor-infiltrating CD8⁺ T cell percentages in both tumor (Fig. 6C) and spleen (Fig. 6F) compartments. Given that the secretion of IFN-γ and granzyme B (GzmB) serves as a functional indicator of cytotoxic T lymphocytes because of their direct tumoricidal effects through tumor cell lysis, we further analyzed the proportion of CD8⁺ T cells coexpressing these effector molecules. Quantitative analysis revealed that LNP@PRO_P+B treatment induced 7.3-, 1.7-, and 2.1-fold higher production of tumor-infiltrating CD8⁺ IFN-γ⁺ T cells compared to PBS, LNP@PRO_P, and LNP@PRO_B groups, respectively (Fig. 6D). Similarly, CD8⁺GzmB⁺ T cell populations in the LNP@PRO_P+B group showed 4.1-, 3.3-, and 1.9-fold increases relative to the three control groups (Fig. 6E). A parallel trend was observed in the spleen (Fig. 6G and fig. S32). Notably, despite the relatively lower baseline proportion of tumor-infiltrating CD8⁺ T cells in the poorly immunogenic 4T1 tumor model compared to B16F10 counterparts (figs. S33 and S34), treatment with LNP@PRO_P+B led to a significant up-regulation of CD8⁺ T cell populations within both tumor and spleen compartments, demonstrating statistically distinct differences from all three control groups. This immunostimulatory effect was similarly reproduced in CT26 tumor–bearing mice, highlighting the translational potential of the therapeutic approach across multiple tumor models. Collectively, these results demonstrate that LNP@PRO_P+B treatment potently stimulates tumor-infiltrating lymphocytes and spleen cells to secrete IFN-γ and GzmB, thereby enhancing tumor cell cytotoxicity and achieving substantial tumor growth suppression. In parallel, NK cells, as a key innate cytotoxic lymphocyte, contribute crucially to antitumor immunity through direct tumoricidal activity and augmentation of adaptive immune responses following cGAS-STING pathway activation. Flow cytometric analysis revealed significant expansion of CD45⁺CD3⁻NK1.1⁺ cell populations in both splenic and tumor tissues of B16F10-bearing mice post–LNP@PRO_P+B treatment (Fig. 6, H to L, and figs. S35 and S36). Notably, the proportions of IFN-γ⁺ and GzmB⁺ NK cells within tumors reached 38.5 and 17.2%, respectively, representing 2.8- and 4.0-fold increases compared to PBS-treated controls (Fig. 6L). Similar up-regulation in NK cell frequencies and effector molecule expression was confirmed in 4T1 and CT26 models (figs. S37 and S38). Emerging evidence highlights the critical regulatory role of tumor-specific effector T cell and regulatory T cell (T_reg cell; CD3⁺CD4⁺CD25⁺Foxp3⁺) balance in determining antitumor immune potency. Subsequent analysis of T_reg populations (gating strategy outlined in fig. S39) demonstrated that LNP@PRO_P+B treatment achieved the most substantial T_reg reduction in tumor microenvironments (Fig. 6, M to O, and figs. S40 and S41), with reduction exceeding 70% compared to PBS-treated controls.

To unequivocally demonstrate the pivotal role of CD8⁺ T cells and NK cells in antitumor immunity, we established a B16F10-Luc melanoma model in C57BL/6 mice and initiated intravenous administration of LNP@PRO_P+B via the tail vein starting on day 7 post–tumor inoculation, with treatment continuing for five consecutive days. To deplete CD8⁺ T cells and NK cells, control groups received intraperitoneal injections of anti-CD8α antibodies (aCD8) and anti-NK1.1 antibodies (aNK1.1) on days 1 and 8 (Fig. 7A). The results revealed that while LNP@PRO_P+B monotherapy exhibited potent tumor suppression (Fig. 7, B and C), its therapeutic efficacy was markedly attenuated when combined with CD8⁺ T cell/NK cell depletion (aCD8 + aNK1.1 + LNP@PRO_P+B). Although residual tumor inhibition remained, likely attributable to the synthetic lethality induced by LNP@PRO_P+B, the absence of CD8⁺ T cells and NK cells compromised long-term tumor control, underscoring their indispensable role in antitumor immunity. To further assess depletion efficiency, we quantified splenic CD8⁺ T cell and NK cell populations at the experimental end point (Fig. 7, D and E). Flow cytometric analysis demonstrated near-complete depletion of both CD8⁺ T cells and NK cells (<1%) in the aCD8 + aNK1.1 + LNP@PRO_P+B treatment group, whereas mice receiving LNP@PRO_P+B alone exhibited significant expansion of these immune effector populations, further corroborating their critical contribution to tumor regression (fig. S42). Collectively, these results establish that LNP@PRO_P+B-mediated STING pathway activation effectively orchestrates a systemic antitumor immune response through coordinated enhancement of cytotoxic lymphocyte and NK cell activity and immunosuppressive cell suppression.

Fig. 7. Therapeutic efficacy of LNP@PRO_P+B in a murine model of lung metastasis.

(A) Schematic illustration of the timeline for B16F10-Luc tumor inoculation and treatments in a C57BL/6 mouse model. ip, intraperitoneally. (B) In vivo bioluminescence imaging of B16F10-Luc tumor growth after administration with different formulations. n = 3 biologically independent samples. d, days. (C) Data analysis of the bioluminescence signals of the mice. Representative flow cytometry analysis plots of CD8⁺ T cells (D) and NK1.1 cells (E) in the spleens of mice after receiving LNP@PRO_P+B and aCD8 + aNK1.1 + LNP@PRO_P+B treatments. (F) Schematic diagram of the treatment process for the lung metastasis model. (G) Representative images of the excised lungs from each treatment group following the completion of the treatment. Yellow dashed lines indicate lung metastases. n = 5 biologically independent samples. (H) H&E-stained sections of the lungs of each treatment group after the end of treatment. (I) Statistics of the number of lung metastatic nodules in each treatment group after the end of treatment. (J) Weight of lung metastatic nodules in each treatment group after the end of treatment. (K) Representative flow cytometry analysis plots of CD8⁺ T cells and CD4⁺ T cells in the spleens of mice after receiving LNP@PRO_P, LNP@PRO_B, and LNP@PRO_P+B treatments. (L) Quantitative analysis of the T_EM cells and T_CM cells in CD8⁺ T cells after the above treatments (n = 5). (M) Representative flow cytometry analysis plots of T_EM cells and T_CM cells in CD8⁺ T cells after the above treatments. (N) Quantitative analysis of the T_EM cells and T_CM cells in CD4⁺ T cells after the above treatments (n = 5). (O) Representative flow cytometry analysis plots of T_EM and T_CM cells in CD4⁺ T cells after the above treatments. Results are presented as the means ± SD.

Systemic immune memory elicited by LNP@PRO_P+B

To investigate whether therapeutic immune activation confers long-term immunologic memory against metastatic dissemination, a pulmonary melanoma metastasis model in immunocompetent C57BL/6 mice through intravenous inoculation of B16F10 tumor cells was established (Fig. 7F). Two weeks postinoculation, animals with confirmed multifocal pulmonary metastases were randomized to receive LNP@PRO_P, LNP@PRO_B, or combination therapy (LNP@PRO_P+B). Quantitative analysis revealed that LNP@PRO_P+B treatment elicited a robust antitumor immune response by significantly inhibiting pulmonary tumor cell proliferation compared to control groups exhibiting extensive metastatic lesions (Fig. 7G and fig. S43). Histopathological evaluation [hematoxylin and eosin (H&E) staining] demonstrated distinct metastatic patterns: lungs from PBS-, LNP@PRO_P-, and LNP@PRO_B-treated mice exhibited widespread neoplastic invasion with significant nuclear pleomorphism, whereas LNP@PRO_P+B treatment resulted in a marked reduction in both the number and size of metastatic foci (Fig. 7H). Quantitative analysis of pulmonary burden further revealed differential therapeutic efficacy across treatment groups. While both LNP@PRO_P and LNP@PRO_B monotherapies exhibited nonsignificant trends toward reduced lung nodule counts compared to PBS controls, the LNP@PRO_P+B combination therapy achieved statistically significant suppression, limiting metastatic nodules to fewer than five per lung (Fig. 7I). Corroborating this finding, lungs from PBS-treated mice demonstrated an 86.9% greater mass compared to those receiving LNP@PRO_P+B, indicative of substantially enhanced metastatic progression in control groups (Fig. 7J). Mechanistic investigation through flow cytometric profiling revealed that LNP@PRO_P+B treatment significantly augmented CD8⁺ T cell infiltration in the spleen, reaching to 1.4 ± 0.11–fold compared to PBS controls (Fig. 7K and fig. S44). Detailed analysis of memory T cell subsets showed that the combination therapy preferentially amplified tumor-specific populations (gating strategy shown in fig. S45): CD8⁺ effector memory T cells (T_EM cells; CD44⁺CD62L⁻) reached 12.0 ± 7.9–fold higher levels than PBS controls (3.1 ± 2.3–fold versus LNP@PRO_P and 6.1 ± 3.6–fold versus LNP@PRO_B), while CD8⁺ central memory T cells (T_CM cells; CD44⁺CD62L⁺) increased to 5.6 ± 2.3–fold compared to PBS baselines (2.1 ± 0.9–fold versus LNP@PRO_P and 4.7 ± 2.0–fold versus LNP@PRO_B). Parallel enhancements were observed in CD4⁺ T_EM or T_CM populations (Fig. 7, L to O). These coordinated immunologic alterations strongly suggest that LNP@PRO_P+B-mediated cGAS-STING pathway activation orchestrates tumor-specific T cell immunity through synthetic lethality mechanisms, effectively containing metastatic dissemination.

To elucidate the antitumor mechanisms underlying LNP@PRO_P+B treatment, we conducted transcriptomic profiling of lung metastases from mice treated with PBS or LNP@PRO_P+B following therapy completion. Differential gene expression analysis, using a fold change threshold >1.5 and adjusted P < 0.05, identified 558 differentially expressed genes (DEGs) in the LNP@PRO_P+B group compared to the control, including of 486 up-regulated and 72 down-regulated genes (Fig. 8, A and B). Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis revealed significant enrichment of DEGs in pathways such as the cytosolic DNA-sensing pathway, NK cell–mediated cytotoxicity, ubiquitin-mediated proteolysis, nuclear factor κB signaling, and cytokine-cytokine receptor interaction (Fig. 8C). Notably, LNP@PRO_P+B demonstrated robust enrichment in the cellular senescence, a pathway canonically activated by genomic instability and DNA damage. This observation mechanistically validates the notion that our therapeutic intervention suppresses the DNA damage response cascade, thereby promoting synthetic lethality in tumor cells. Gene Ontology (GO) analysis further indicated that LNP@PRO_P+B treatment enhanced biological processes including “positive regulation of T cell–mediated immunity,” “DC antigen processing and presentation,” and “regulation of type I interferon–mediated signaling pathway” (Fig. 8D). Gene set enrichment analysis of lung metastases from LNP@PRO_P+B-treated mice showed up-regulation of immune activation–related pathways and suppression of DNA damage repair pathways such as base excision repair, nucleotide excision repair, and HR, confirming that LNP@PRO_P+B potently inhibits DNA damage repair processes (Fig. 8E). Collectively, these transcriptomic analyses elucidate that LNP@PRO_P+B exerts antitumor effects through a dual mechanism: enhancing immune surveillance via DC/T cell/NK cell–mediated responses and concurrently impairing DNA repair pathways, thereby reinforcing tumor cell vulnerability. Together, these findings provide mechanistic insights into the multifaceted antitumor activity of LNP@PRO_P+B in metastatic lung lesions.

Fig. 8. Transcriptomic analysis of lung metastatic following LNP@PRO_P+B treatment versus PBS control.

(A) Volcano plot illustrating DEGs between PBS and LNP@PRO_P+B groups, with significantly up-regulated genes shown in red and down-regulated genes in blue. (B) Heatmaps display the log₂ fold change in gene expression for LNP@PRO_P+B versus PBS treatment (n = 3 for each group). (C) Enriched pathways of KEGG gene sets in LNP@PRO_P+B-treated tumor in the lung. (D) GO enrichment evaluation of the pathways in the LNP@PRO_P+B versus PBS groups. (E) KEGG database gene set enrichment analysis enrichment ridge map. Enrichment score (ES) > 0 indicates that the pathway is activated and the enriched core genes are up-regulated genes, while ES < 0 indicates that the pathway is inhibited and the enriched core genes are down-regulated genes.

DISCUSSION

The activation of the STING pathway plays a central role in antitumor immunity. Here, we report a promising therapeutic strategy: PROTAC-mediated degradation of PARP1 and BRD4 induces synthetic lethality, increases cytoplasmic DNA accumulation, and activates the STING pathway–driven innate immune cascade. In vitro and in vivo experiments demonstrate that PRO_P+B potently induces DNA damage, evidenced by marked up-regulation of γ-H2AX. Sustained DNA damage—resulting from the loss of DNA repair responses because of PARP1/BRD4 degradation—promotes the release of dsDNA from the nucleus to the cytoplasm. Accumulation of cytoplasmic DNA eventually surpasses the activation threshold of the STING pathway, leading to its local activation. Notably, this approach exhibits broad applicability: PRO_P+B elicits synthetic lethality across multiple tumor cell lines, with CI values <1 in all treated groups, indicating synergistic cytotoxicity between PRO_P and PRO_B. Furthermore, in three tumor models with distinct immunogenicity profiles, LNP@PRO_P+B demonstrates robust antitumor efficacy: DNA released from dying tumor cells activates the cGAS-STING pathway in adjacent immune cells, such as DCs, driving increased T cell infiltration, DC maturation, and NK cell enrichment. This strategic integration of synthetic lethality with immunogenic stress responses establishes a previously unidentified paradigm for potentiating cancer immunotherapy and expanding the pool of treatable patients.

MATERIALS AND METHODS

Compounds

The synthesis of products and their respective intermediates are detailed in the Supplementary Materials.

Cell culture

B16F10, 4T1, H22, KPC, and CT26 cell lines were obtained from American Type Culture Collection. DC2.4 and THP-1 cell lines were obtained from Millipore-Sigma. B16F10, 4T1, H22, KPC, and CT26 cells were cultured in RPMI 1640 (Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco). DC2.4 cells were cultured with RPMI 1640 supplemented with 10% FBS, 1× nonessential amino acids (Gibco), 1× Hepes buffer (Gibco), and 0.0054× β-mercaptoethanol (Gibco). THP-1 cells were cultured with RPMI 1640 supplemented with 10% FBS and 0.05 mM 2-mercaptoethanol. All the adherent cell lines were cultured at 37°C and 5% CO₂ in a humidified atmosphere.

Experimental mouse models

All animal procedures were approved by the Institutional Animal Care and Use Committee of South China University of Technology (protocol no. 2023066). Female C57BL/6J, BALB/cJ, and NCG mice (4-week-old) were housed under controlled conditions: 12-hour light/dark cycle, ambient temperature of 20° to 22°C, humidity of 45 to 65%, and noise <50 dBA.

For tumor-bearing models, mice were randomly assigned to treatment groups in a blinded manner. Upon tumor reaching ~70 mm³, mice were administered intravenous injections of PBS, LNP@PRO_P, LNP@PRO_B, or LNP@PRO_P+B. Survival end points were defined as the tumor volume exceeding 2000 mm³ or body weight loss >30%, at which point mice were euthanized.

WB analysis

To evaluate the degradation of PARP1 and BRD4, cells were seeded in six-well plates. At ~80% confluency, cells were treated with PRO_P (0.01 to 25 μM), PRO_B (0.01 to 25 μM), or their combination (PRO_P+B) for 12 or 24 hours. After PBS washing, cells were lysed with radioimmunoprecipitation assay buffer containing protease inhibitors. Protein concentrations were determined using a bicinchoninic acid assay kit (Pierce). Equal amounts of lysates (20 μg per lane) were separated on 8% SDS–polyacrylamide gel electrophoresis gels and transferred to polyvinylidene difluoride membranes. Membranes were blocked with 5% bovine serum albumin in tris-buffered saline with Tween 20 for 2 hours at room temperature (RT), followed by overnight incubation at 4°C with primary antibodies: anti-PARP1 (1:1000; Abcam, ab1912), anti-BRD4 [1:1000; Cell Signaling Technology (CST), no. 13440], and anti-GAPDH (glyceraldehyde-3-phosphate dehydrogenase; 1:5000; Proteintech, 60004-1-Ig). After washing with tris-buffered saline with Tween 20, membranes were incubated with HRP (horseradish peroxidase)–conjugated secondary antibodies (1:5000; Jackson ImmunoResearch) for 2 hours and developed using an enhanced chemiluminescence substrate (Millipore).

To evaluate STING signaling, B16F10, 4T1, H22, CT26, and KPC tumor cells were treated with PRO_P (1 μM), PRO_B (1 μM), or PRO_P+B for 24 hours. Conditioned media (50%, v/v) from treated tumor cells, supplemented with fresh medium, were applied to DC2.4 cells for 48 hours. WB was performed as above using antibodies against cGAS (1:1000; CST, no. 15102), p-STING (Ser³⁶⁶: 1:500; CST, no. 72971), and p-IRF3 (Ser³⁹⁶: 1:500; CST, no. 4947).

Immunofluorescence staining

Degradation of BRD4 and PARP1 was confirmed through confocal laser scanning microscopy (CLSM). Tumor cells were seeded in six-well plates at 5 × 10⁵ cells per well for 24 hours before treatment with PRO_P (1 μM), PRO_B (1 μM), or their combination (PRO_P+B) for 24 hours. Cells were subsequently harvested, washed thrice with PBS, and fixed with 4% paraformaldehyde for 15 min at RT. For CLSM detection, fixed cells were incubated with a rabbit anti-BRD4 or anti-PARP1 primary antibody at 37°C for 1 hour, followed by three PBS washes and incubation with an Alexa Fluor 488– or Alexa Fluor 647–conjugated goat anti-rabbit secondary antibody (1:500; Invitrogen) under light-protected conditions at 37°C for 1 hour. Nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI; 5 μg ml⁻¹, 15 min) before imaging with a Zeiss LSM 880 system. For quantitative assessment, the fluorescence intensity was analyzed using ZEN Microscopy Software.

Detection of dsDNA in the cytoplasm

The cells were treated with PRO_P (1 μM), PRO_B (1 μM), or their combination (PRO_P+B) for 24 hours. Then, the cells were fixed with 4% paraformaldehyde, selective plasma membrane permeabilization was then performed by incubating the fixed cells with 0.02% saponin/PBS (5 min), and then the cells were blocked with 2.5% normal goat serum. After overnight incubation with an anti-dsDNA antibody (AE-2; Sigma-Aldrich, MAB1293; 1:200) at 4°C, cells were stained with an Alexa Fluor 488–conjugated secondary antibody (Thermo Fisher Scientific, A-11032; 1:400). For quantitative analysis, cytoplasmic fractions were isolated by differential centrifugation (500g, 10 min) following liquid nitrogen freeze-thaw cycles. The genomic DNA extraction reagent kit (Tiangang, A0605A) was used to extract dsDNA, and the extracted dsDNA was quantitatively analyzed using a NanoDrop 2000c spectrophotometer (Thermo Fisher Scientific).

Down-regulation of PD-L1 expression by PRO_P+B

PD-L1 degradation was confirmed through integrated flow cytometry, CLSM, and WB analyses. Tumor cells were seeded in six-well plates at 5 × 10⁵ cells per well for 24 hours before treatment with PRO_P (1 μM), PRO_B (1 μM), or their combination (PRO_P+B) for 12 hours. For CLSM detection, fixed cells were incubated with a rabbit anti–PD-L1 primary antibody (1:200; CST, no. 13684) at 37°C for 1 hour. For quantitative assessment, parallel samples were processed for WB using 20 μg of protein lysates probed with a PD-L1–specific antibody (1:1000; Proteintech, 17952-1-AP). Furthermore, parallel samples were processed for flow cytometry analysis (BD FACS Celesta, 10,000 events per sample) and then specifically visualized as a heatmap generated using GraphPad Prism 9.0.

Comet assay for DNA damage

Following 12-hour treatment with PRO_P (1 μM), PRO_B (1 μM), or PRO_P+B combination, cells were trypsinized and resuspended in PBS. Cell suspensions were mixed with low-melting-point agarose [0.7% (w/v) in PBS, 37°C] and layered onto precoated slides (1% normal-melting-point agarose). After solidification (4°C, 15 min), slides were immersed in lysis buffer (2.5 M NaCl, 100 mM EDTA, 10 mM tris-HCl, and 1% Triton X-100, pH 10) for 1 hour at 4°C. Electrophoresis was conducted in alkaline buffer (300 mM NaOH and 1 mM EDTA, pH > 13) at 20 V for 20 min. DNA was stained with propidium iodide (2.5 μg ml⁻¹), and comet formation was analyzed using fluorescence microscopy.

Immunofluorescence of γ-H2AX

Posttreatment cells were seeded on 12-mm glass coverslips for immunofluorescence processing. Following fixation with 4% paraformaldehyde in PBS (15 min, RT), cells underwent sequential permeabilization with 0.5% Triton X-100 (10 min) and blocking in PBS containing 3% goat serum (Sigma-Aldrich, G9023), bovine serum albumin (1 mg ml⁻¹; Sigma-Aldrich, A7906), 0.1% Triton X-100, and 1 mM EDTA (30 min). γ-H2AX foci were detected using the DNA Damage Assay Kit (Beyotime, C1063S) per the manufacturer’s protocol: Primary antibody incubation (1:200 in blocking buffer, 1 hour, RT) was followed by incubation with an Alexa Fluor 488–conjugated secondary antibody (Invitrogen, A-11008; 1:500; 1 hour, RT). Nuclei were stained with DAPI (1 μg ml⁻¹, 5 min). Imaging was performed on a Zeiss LSM880 confocal microscope using 405-nm (DAPI) and 488-nm (Alexa Fluor 488) excitation lasers.

ELISA of cGAMP and IFN-β release

The tumor cells were first treated with various formulations according to the procedure described above. Tumor cell–conditioned media were prepared by combining 50% (v/v) clarified supernatants (obtained by centrifugation at 500g for 10 min) with fresh complete medium. These media were applied to C57BL/6 BMDCs for 24 hours of stimulation. cGAMP and IFN-β levels in tumor/BMDC supernatants were quantified using commercial ELISA kits (VeriKine Mouse IFN-β Kit, PBL Assay Science, cGAMP ELISA Kit, MyBioSource) following the standardized protocol: After loading standards/samples (100 μl per well) into antibody-precoated plates, a 90-min incubation at 37°C preceded five cycles of washing with PBS and 0.05% Tween 20. Biotinylated detection antibodies in assay buffer were subsequently incubated (1:1000 dilution, 60 min, 37°C) followed by streptavidin-HRP conjugate treatment (1:2000, 30 min, 37°C). Following final washes, a trimethylboron substrate (BD Biosciences) was developed for 15 min at 37°C before reaction termination with 1 MH₂SO₄. Absorbance was measured at 450 nm (reference, 570 nm) using a Multiskan GO microplate reader (Thermo Fisher Scientific), with sample concentrations calculated against standard curves.

BMDC and BMDM isolation and maturation

Femurs and tibias from female wild-type C57BL/6 mice (8 to 12 weeks) were aseptically dissected and flushed with ice-cold PBS (Gibco, 10010023) using 25G needles to harvest bone marrow progenitors. Following erythrocyte lysis with ACK (ammonium-chloride-potassium) buffer (Lonza, 10-548E), a portion of the cells were seeded at 2 × 10⁶ cells per 100-mm non–tissue culture–treated petri dish (Corning, 430591) in RPMI 1640 medium supplemented with 10% heat-inactivated FBS, 1% penicillin/streptomycin (Sigma-Aldrich, P4333), and recombinant murine granulocyte-macrophage colony-stimulating factor (GM-CSF; 20 ng ml⁻¹). On day 3 postseeding, the unattached cells were used as BMDCs, and cultures received an additional 10-ml complete medium containing GM-CSF (40 ng ml⁻¹) to maintain the final cytokine concentration. From day 6 onward, 50% medium exchange was performed every 48 hours by centrifuging conditioned media (300g, 4°C, 10 min) and replenishing with fresh GM-CSF–containing medium. Another part of the cells was collected and cultured in complete BMDM medium [complete α-minimum essential medium supplemented with macrophage colony-stimulating factor (50 ng ml⁻¹)]. Three days later, the adherent cells were used as BMDMs.

BMDCs or BMDMs were seeded in 24-well plates at 2 × 10⁵ cells per well and incubated with the tumor cells that were pretreated with PRO_P (1 μM), PRO_B (1 μM), or PRO_P+B. After 24 hours, the supernatant was removed, and the cells were collected, washed, stained, fixed, and analyzed by flow cytometry. The following antibodies were used: FITC-CD11c, PE-CD11b, APC-CD86, PE-CD80, and FITC-CD86.

In vitro cytotoxicity

Cytotoxic effects were evaluated using the Cell Counting Kit-8 assay. Tumor cells were seeded in 96-well plates at 5 × 10³ cells per well and cultured for 24 hours. Cells were then exposed to the following: (i) LNP@PRO_P (0.01 to 25 μM), (ii) LNP@PRO_B (0.01 to 25 μM), or (iii) their combination (LNP@PRO_P+B) for 24 hours. Following PBS washing (3 × 100 μl per well), 100 μl of serum-free medium containing 10% Cell Counting Kit-8 reagent was added to each well and incubated under light-protected conditions (37°C, 4 hours). Absorbance was measured at a 450-nm reference using a Multiskan GO microplate reader (Thermo Fisher Scientific). Cell viability was calculated as (OD_treatment/OD_control) × 100%, with IC₅₀ (median inhibitory concentration) values determined through nonlinear regression analysis (four-parameter logistic model) in GraphPad Prism 9.3.

Clonogenic assay

Single-cell suspensions were prepared through 40-μm-mesh filtration (Corning) and plated in six-well plates at 1000 cells per well for overnight adherence. Following 12 hours of treatment with LNP@PRO_P (1 μM), LNP@PRO_B (1 μM), or LNP@PRO_P+B combination, cells underwent 7-day culture in complete medium. Colonies were stained with 0.5% crystal violet in 20% methanol (30 min, RT). Stained colonies were quantified using a calibrated flatbed scanner (Epson Perfection V600) with a 1200–dpi (dots per inch) resolution.

In vivo anticancer study

For the B16F10 melanoma model, female C57BL/6 mice (6 to 8 weeks) or NCG mice (4 weeks) received subcutaneous injections of 5 × 10⁵ B16F10 cells in 100 μl of PBS (right flank) on day 0. From days 1 to 5 postinoculation, tumor-bearing mice were administered intravenous formulations containing the following: (i) LNP@PRO_P (10 mg kg⁻¹), (ii) LNP@PRO_B (2 mg kg⁻¹), and (iii) LNP@PRO_P+B (n = 7 per group). For the depletion of CD8⁺ T cells and NK1.1 cells in vivo, anti-CD8a antibodies (aCD8, 300 μg per mouse, BioXcell, catalog number BE0061, clone number 2.43) and anti-NK1.1 antibodies (aNK1.1, 300 μg per mouse, BioXcell, catalog number BE0036, clone number PK136) were intraperitoneally injected.

For the CT26 colon carcinoma model, female BALB/c mice (4 to 6 weeks) were subcutaneously implanted with 5 × 10⁵ CT26 cells (right flank) in 100 μl of PBS. Identical dosing regimens as the B16F10 model were administered on days 1 to 5.

For the orthotopic 4T1 breast cancer model, female BALB/c mice (6 weeks) underwent mammary fat pad inoculation with 5 × 10⁵ 4T1 cells (100 μl of PBS). Identical dosing regimens as the B16F10 model were administered on days 1 to 5. For tumor monitoring, all models used bidimensional caliper measurements (tumor volume = 0.5 × L × W²) every 48 hours until the end point (2000 mm³).

Lung metastasis model

For the B16F10 lung metastasis model, C57BL/6 mice (4 to 6 weeks, female) were injected intravenously with 2 × 10⁵ B16F10 cells. Two weeks later, the mice developed multifocal metastases on both lungs. Tumor-bearing mice were administered intravenous formulations containing the following: (i) LNP@PRO_P (10 mg kg⁻¹), (ii) LNP@PRO_B (2 mg kg⁻¹), and (iii) LNP@PRO_P+B (n = 7 per group). After treatment, the mice were euthanized, and the lung tissues were harvested for photography. The pulmonary metastatic nodules were counted manually. The tissues were stained with H&E to assess toxicity or lung metastasis.

Flow cytometry assay

After completion of treatment, mice bearing B16F10/CT26/4T1 tumors were euthanized, and tumor-draining lymph nodes, spleens, and tumors were collected and prepared into single-cell suspensions. Briefly, tumor-draining lymph nodes and spleens were gently ground in a PBA solution (PBS supplemented with 5% BSA) and filtered through 200-mesh nylon. Furthermore, the red blood cells in a single-cell suspension of spleens were removed with 1× Red Blood Cell Lysis Buffer (Solarbio), the digestion was terminated by addition of PBA solution, and the red blood cells were centrifuged and resuspended with a PBA solution. Tumor tissues were cut into pieces and placed into 2% FBS RPMI digestion solutions containing collagenase IV, hyaluronidase, and DNase for 30 min. Subsequently, the digestion solutions were filtered through 200-mesh nylon. Then, the digestion solutions were centrifuged at 2000 rpm for 5 min, the supernatant was removed, and the precipitate was resuspended in 40% Percoll. The precipitate was resuspended in 1× Red Blood Cell Lysis Buffer, the digestion was terminated by addition of PBA solution, and the precipitate was centrifuged and resuspended with a PBA solution to obtain a tumor tissue cell suspension. Cells were blocked with anti-mouse CD16/32 for 20 min on ice.

For DC maturation analysis, the draining lymph node single-cell suspension was stained with anti–CD45-APC/Cy7, anti–CD11c-FITC, anti–CD80-PE, and anti–CD86-APC antibodies. Last, the cells were analyzed with flow cytometry.

For the analysis of the lymphocytes in the spleen, the spleen single-cell suspension was stained with anti–CD45-APC/Cy7, anti–CD3-FITC, anti–CD4-BV510, and anti–CD8-BV785 antibodies. Last, the cells were analyzed with flow cytometry.

For the analysis of lymphocytes in the tumor analysis, the tumor single cells were stained with a fluorescently labeled antibody according to the manufacturer’s protocols. To examine intratumoral infiltrating T_reg cells (CD45⁺CD3⁺CD4⁺CD25⁺Foxp3⁺), the T lymphocytes were stained with anti–CD45-APC/Cy7, anti–CD3-FITC, anti–CD4-BV510, anti–CD25-APC, and anti–Forxp3-PE antibodies according to the manufacturer’s protocols. To investigate CD8⁺ (CD45⁺CD3⁺CD8⁺) T lymphocytes, the total T lymphocytes were stained with anti–CD45-APC/Cy7, anti–CD3-FITC, and anti–CD8-BV785 antibodies according to the manufacturer’s protocols and analyzed with flow cytometry.

Statistics and reproducibility

All measurements were performed with three or more independent replicates from separate experiments. The exact sample size and statistical test for each experiment are described in the relevant figure legends. All results are presented as the means ± SEM. Statistical analyses were conducted using GraphPad Prism with Student’s t test, a one-way analysis of variance (ANOVA), or a two-way ANOVA.

Supplementary Materials

This PDF file includes:

Supplementary text

Scheme S1

Figs. S1 to S45