PD-L1-targeted photodynamic therapy orchestrates checkpoint blockade and immunogenic cell death for synergistic cancer immunotherapy

Abstract

Inhibiting the PD1/PD-L1 interaction is crucial for developing novel cancer immunotherapies, particularly to reduce systemic toxicity and enhance patient response rates. In this study, we designed and synthesized Photodegradation-Targeting Chimeras (PDTACs) by conjugating a clinically approved photosensitizer, verteporfin, to a PD-L1-targeted peptide. Our optimized chimera, PPA-VPF, demonstrates a dual mechanism of action in cancer immunotherapy, resulting from singlet oxygen generated under light irradiation. The proximity-generated singlet oxygen effectively degrades PD-L1 in cancer cells through immediate protein breakdown and resulted in subsequent lysosomal-dependent degradation hours after irradiation. Additionally, the non-proximity-generated singlet oxygen induces immunogenic cell death (ICD) through cytotoxic effects. In mouse models with immune cold tumors, PPA-VPF elicited robust adaptive antitumor immunity and effectively inhibited the growth of both primary and distant tumors. This PD-L1-targeted PDTAC achieved immune checkpoint blockade and ICD induction in a single therapeutic mode using one molecular species, presenting a novel strategy for combinational immunotherapy, particularly in immune cold tumors.

Graphical abstract

1. Introduction

Immunotherapies, particularly immune checkpoint blockade (ICB), have recently revolutionized cancer treatment by enhancing the host's immune system [[1], [2], [3]]. Notably, the inhibition of the PD1/PD-L1 (programmed cell death protein 1/programmed death-ligand 1) interaction has achieved significant success in treating various cancers [[4], [5], [6]]. While thousands of clinical trials based on PD1/PD-L1 inhibition are currently ongoing worldwide, several monoclonal antibodies have already received approval from the U.S. Food and Drug Administration [7]. Despite their considerable efficacy, only a limited number of patients benefit from mono-ICB therapy. Cold tumors, characterized by low activity of suppressor immune cells, often exhibit poor therapeutic responses to ICBs [8,9]. Consequently, various combination strategies have been extensively explored to enhance the efficacy of ICB therapy, both in preclinical studies and clinical trials [[10], [11], [12], [13], [14], [15]]. In fact, more than 80% of active PD1/PD-L1 trials are testing combination strategies with other therapies, including chemotherapies and radiotherapies [7]. Certain cytotoxic drugs, such as oxaliplatin, can enhance tumor immunogenicity by inducing immunogenic cell death (ICD) [16,17]. These dying tumor cells release tumor-specific antigens and damage-associated molecular patterns (DAMPs), which attract and activate dendritic cells (DCs) [18]. As a result, DCs stimulate the activation of effector T cells through antigen presentation, leading to a long-lasting antitumor immune response. However, introducing additional therapeutic modalities in combination therapy raises potential safety concerns that may restrict clinical translation.

Additionally, autoimmune-mediated side effects have emerged as a safety concern regarding PD-1/PD-L1 inhibition [19,20]. These immune-related adverse events (irAEs) encompass various endocrine dysfunctions, pneumonitis, hepatitis, and colitis. Due to the expression of PD-L1 in some normal tissues, these on-target side effects are challenging to prevent when therapeutic antibodies are administered systemically. Thus, there is an urgent need for selective inhibition of PD1/PD-L1 specifically in cancer cells to reduce irAEs in patients.

We previously developed a targeted protein photolysis approach, known as the photodegradation-targeting chimera (PDTAC) strategy, to degrade the cytosolic protein GPX4 [21]. These chimeras consist of three functional modules: a photosensitizer, a protein-targeted ligand, and a linker to conjugate the two parts. Under light irradiation, the photosensitizer generates reactive oxygen species (ROS), particularly singlet oxygen (¹O2). The targeting ligand directs most of the photoinduced ¹O2 to the protein of interest (POI). The highly reactive ¹O2 can inactivate or even degrade the POI by oxidizing amino acids or breaking peptide backbones [[22], [23], [24], [25]]. Compared to other targeted protein degradation methods [[26], [27], [28]], our PDTAC strategy retains the high spatiotemporal precision of light irradiation and is expected to have broad applicability to membrane proteins, including PD-L1. Furthermore, if the targeting ligand has a mild or negligible inhibitory effect on the POI [21], PDTAC may provide an additional advantage by mitigating on-target side effects.

In addition to the targeted degradation of proteins using proximity-generated ROS, non-proximity-generated ROS typically react indiscriminately with biomolecules, including various proteins [[29], [30], [31]]. Sufficient levels of ROS can directly kill cancer cells due to accumulated damage to various cellular molecules, which is the primary mechanism of action of photodynamic therapy (PDT) using conventional photosensitizers. This form of cell death is often immunogenic, making PDT a treatment modality that triggers ICD, much like cytotoxic drugs [32,33]. Due to its noninvasiveness and effectiveness against certain chemo-resistant cancers, PDT-induced immunotherapy has emerged as a promising option for combination therapy with PD1/PD-L1 inhibition [[34], [35], [36]].

In this study, we designed PDTACs to synergistically degrade PD-L1 and induce ICD (Scheme 1). A D-peptide with good metabolic stability and strong affinity toward PD-L1 was chosen as the targeting ligand. Based on binding affinity optimization, two water-soluble PEG groups were utilized as the linker. This chimera demonstrated selectivity toward PD-L1-positive cells. As anticipated, we observed dual immunological effects from the singlet oxygen generated under light irradiation. The proximity-generated singlet oxygen degraded PD-L1 to block the immune checkpoint, while the non-proximity-generated singlet oxygen triggered ICD to activate DCs. In mouse xenograft models featuring immune cold tumors, light treatment following intravenous injection of the chimera effectively elicited potent antitumor immunity and inhibited the growth of both primary and distant tumors. Therefore, our PDTAC incorporates the synergistic effects of both ICB and ICD induction for combinatorial immunotherapy. Additionally, its selectivity for PD-L1-positive cells, along with the precision of light irradiation, endows our chimeras with high biocompatibility and biosafety.

Scheme 1.

Schematic illustration of the PD-L1-targeted photosensitizer chimera and its dual mechanism of action in cancer photoimmunotherapy. (a) Schematic structure of the PDTAC molecule. (b) Principle of singlet oxygen generated from photosensitizer. (c) Synergistic effects of PD-L1-drgradation and ICD-induction in cancer photoimmunotherapy.

2. Results and discussion

2.1. Synthesis and characterization of PPA-VPF chimeras

We designed four PDTAC molecules aimed at degrading PD-L1, as illustrated in Fig. 1b. The targeting ligand is a D-peptide, ^DN^DY^DS^DK^DP^DT^DD^DR^DQ^DY^DH^DF (termed PPA), which demonstrates a binding affinity of 0.38 μM and improved proteolytic stability in serum [37]. Verteporfin (VPF), an FDA-approved photosensitizer, exhibits an excellent ability to generate singlet oxygen and has strong absorption in the red-light region at 689 nm [38]. These two moieties were conjugated using a linker, which was either a hydrophobic C6 or a hydrophilic PEG. We selected various PEG lengths, specifically PEG2, PEG5, and PEG10, to optimize the linker length. The four chimeras were synthesized via solid-phase peptide synthesis (SPPS) and were designated as PPA-VPF, PPA-VPF-1, PPA-VPF-2, and PPA-VPF-3 (see Fig. 1a–Table S1, Fig. S1).

Fig. 1.

Synthesis and characterization of the PDTAC molecules. (a) Synthesis of PDTACs by SPPS. (b) Summary of the binding affinities toward PD-L1 measured by SPS. (c) Measurements of the binding affinity of PPA-VPF, PPA and VPF toward PD-L1 using SPR. (d) UV-Vis absorption spectra of PPA-VPF and VPF and in PBS or PBS/ACN (1:1). (e) Fluorescence spectra of PPA-VPF and VPF in PBS or PBS/ACN (1:1). (f). Amount of singlet oxygen generated from VPF and PPA-VPF (20 μM) in PBS or PBS/ACN (1:1) measured by EPR using 4-hydroxy-2,2,6,6-tetramethylpiperidine (4-OH-TEMP, 200 mM) as the spin trap. Light irradiation was performed using a 300 W Xenon arc lamp with a 600 nm bandpass filter (∼1.5 mW/cm²) for the designated time.

After purification by high-performance liquid chromatography (HPLC) (Fig. S8–12), we examined the morphologies of the chimeras in phosphate-buffered saline (PBS) using dynamic light scattering (DLS) (Table S2). The chimeras with hydrophilic linkers, along with the parental PPA, exhibited hydrodynamic sizes ranging from 100 to 300 nm. The chimeras formed a loose structure due to the reasonable solubility of PPA itself, as indicated by the high polydispersity index (PDI, ∼0.5). Consistently, transmission electron microscopy (TEM) further confirmed the loose nanostructure of PPA-VPF and the tendency to form larger particles with irregular morphology (Fig. S2d). In contrast, PPA-VPF3, which contains a hydrophobic C6 linker, had a hydrodynamic size greater than 1000 nm. Additionally, the zeta potentials of these chimeras ranged from +2.3 to +4.8 mV in PBS (pH 7.4), comparable to that of PPA (+2 mV).

To achieve targeted photodegradation, it is critical for PDTACs to bind to PD-L1. We employed surface plasmon resonance (SPR) spectroscopy to measure their binding affinities toward human PD-L1 immobilized on a sensor chip (Fig. 1b and c, Fig. S2). First, VPF itself exhibited no specific binding to PD-L1. Second, all PDTACs with hydrophilic linkers demonstrated reasonable affinities comparable to that of their parental peptide PPA, with the K_D increasing slightly with longer linkers. In particular, the K_D of PPA-VPF with PEG2 remained in the submicromolar range. Third, PPA-VPF-3 with the hydrophobic linker did not show significant binding to PD-L1. Therefore, both VPF and the short PEG linker had a negligible impact on the binding between PPA and PD-L1, suggesting that the loosely assembled structure formed by these chimeras could break down upon exposure to PD-L1.

Based on the SPR and DLS measurements, we selected PPA-VPF as the PDTAC for PD-L1 degradation. The shortest linker of PPA-VPF may also reduce the diffusion distance of photoinduced ¹O₂ from VPF to PD-L1. In PBS/ACN (1:1) solutions, both PPA-VPF and VPF exhibited identical ultraviolet-visible (UV-Vis) and fluorescence spectra (Fig. 1d and e). Notably, the amount of ¹O₂ generated by PPA-VPF upon red light irradiation was comparable to that produced by VPF alone (Fig. 1f). These results demonstrate that the conjugation strategy with a linker does not disrupt the electronic structure of VPF. In PBS, PPA-VPF displayed a UV-Vis spectrum that was slightly different from that of VPF (Fig. 1d). Furthermore, the fluorescence intensity and the amount of photoinduced ¹O₂ in PPA-VPF were lower than those in VPF (Fig. 1e and f). These findings align with the loosely assembled structure of PPA-VPF revealed by DLS in PBS (Table S2), which may lead to fluorescent quenching of VPF and attenuate O₂ exposure.

2.2. PPA-VPF selectively targets cancer cells with high PD-L1 expression

Tumor-targeting photosensitizers are an effective strategy in PDT to enhance their accumulation in tumors [35]. Since PD-L1 is often overexpressed in cancers, PPA has been previously shown to accumulate in mouse xenografts [37]. Therefore, we investigated whether PPA-VPF could selectively target cancer cells with high PD-L1 expression. Using the intrinsic fluorescence of VPF, we analyzed the subcellular distribution of PPA-VPF through confocal microscopy. The imaging revealed distinct subcellular distribution patterns of PPA-VPF that depended on PD-L1 expression status (Fig. 2a). In wild-type mouse colon cancer MC38 cells, PPA-VPF exhibited significantly higher intracellular accumulation compared to PD-L1 knockout counterparts after 8 h of incubation. Furthermore, the breast cancer cell lines MDA-MB-231 and MCF-7 were selected as PD-L1-high and PD-L1-low models, respectively. Confocal imaging indicated that more PPA-VPF was enriched in MDA-MB-231 cells after 8 h of incubation (Fig. 2b). Consistently, PD-L1-high MDA-MB-231 cells displayed greater adherence of PPA-VPF than MCF-7 cells at all three tested concentrations in flow cytometry (Fig. 2c).

Fig. 2.

Targeting of PPA-VPF to PD-L1 in cancer cells. (a) Distribution of PPA-VPF or VPF in MC38 WT and PD-L1 KO cells revealed by confocal microscopy. Western blotting confirms PD-L1 knockout efficiency with β-actin as a loading control. (b) PD-L1 expression (green) and distribution of PPA-VPA or VPF (red) in MDA-MB-231 cells and MCF-7 cells at a concentration of 1 μM (incubated for 8 h). (c) Cellular adhesion of PPA-VPF and VPF in two breast cancer cell lines. (d) Time dependence of the cellular uptake of PPA-VPF and VPF in MDA-MB-231 cells. Scale bars represent 20 μm.

To elucidate the cellular uptake mechanism of PPA-VPF, we applied various endocytic inhibitors in MDA-MB-231 cells (Fig. S3a). The cellular uptake of PPA-VPF was partially inhibited by MβCD (an inhibitor of caveolae-mediated endocytosis) but not by chlorpromazine (CPZ, an inhibitor of clathrin-mediated endocytosis) or colchicine (an inhibitor of macropinocytosis). These results indicate that the primary mechanism of cellular uptake for PPA-VPF is mediated by endocytosis through PD-L1 receptors.

We further assessed the contribution of VPF to this selectivity over PD-L1. Under the same conditions, VPF uptake was similar between the two cell lines, as evidenced by both confocal imaging and flow cytometry (Fig. 2b and c). Time-dependent measurements also demonstrated that VPF was taken up more rapidly than PPA-VPF (Fig. 2d). While VPF uptake increased with incubation time over 16 h, PPA-VPF uptake reached saturation after 8 h.

2.3. PPA-VPF photodegrades PD-L1 through a dual mechanism

To investigate the photodegradation effect of PPA-VPF, we analyzed the changes in protein abundance of PD-L1 in both MC38 cells and MDA-MB-231 cells. Treatment with PPA-VPF significantly decreased PD-L1 expression levels measured by western blotting (Fig. 3a, Fig. S4a–b). Flow cytometry analysis also showed that the photodegradation of PD-L1 depended on the dose of PPA-VPF (Fig. 3b). Consistently, confocal microscopy revealed a clear decrease in fluorescence intensity of PD-L1 in PPA-VPF-treated cells (Fig. 3d). Further experiments revealed that PPA-VPF selectively degraded PD-L1 without affecting PD-1 (Fig. S4c–d). Overall, these results indicate that PPA-VPF can effectively photodegrade PD-L1 in cancer cells. In contrast, treatments with PPA, VPF, or their 1:1 mixture (PPA + VPF) had a negligible effect on PD-L1 abundance (Fig. 3b–d), demonstrating that the conjugation of PPA and VPF is essential for PD-L1 degradation.

Fig. 3.

In vitro photodegradation of PD-L1 in cancer cells. Analysis of PD-L1 abundance by western blotting (a), flow cytometry (b, c) and confocal imaging (d) in MDA-MB-231 cells. (e) Time dependence of PD-L1 degradation after PPA-VPF treatment and the blocking effect of chloroquine. (f) Coomassie blue staining of PD-L1 photolyzed in mixed solution (concentrations of PPA, PPA-VPF and VPF were 10 μM). (g) Time dependence of cellular PD-L1 distribution (green) after incubation with PPA-VPF (red). (h) Two mechanisms by which PPA-VPF degrades PD-L1 in cells. Irradiation was performed using a 300 W Xenon arc lamp (600 nm bandpass filter, 1.5 mW/cm²) for the designated time. Scale bars represent 20 μm.

We also tracked changes in PD-L1 abundance over time after light irradiation in cancer cells (Fig. 3e). A decrease in PD-L1 abundance was observed as early as 1 h after irradiation, and interestingly, this decline continued even 8 h post-irradiation. To investigate the degradation mechanism of PPA-VPF treatment, we employed the Bradford protein assay with Coomassie blue to monitor the photolysis process in PD-L1 solutions (Fig. 3f). The parental peptide PPA showed no effect on PD-L1 in SDS-PAGE analysis under light irradiation, indicating that neither PPA nor light could degrade PD-L1. In contrast, significant reductions in PD-L1 were observed after 10 or 20 min of light irradiation with PPA-VPF. As expected, VPF alone had only a minimal effect on PD-L1 upon irradiation.

Furthermore, l-ascorbic acid (1 mM) was used to scavenge ROS in MDA-MB-231 cells during irradiation. l-ascorbic acid significantly mitigated PD-L1 degradation in PPA-VPF-treated cells exposed to irradiation (Fig. S4e), confirming that photo-induced ROS is crucial for the early PD-L1 degradation. Consistent with our previous findings on the photodegradation of GPX4 [21], these results further support the notion that the targeting ligand directs photoinduced singlet oxygen (¹O₂) to effectively degrade the protein of interest. Therefore, the early decrease in PD-L1 abundance after light irradiation can be attributed to the immediate photo-breakdown effect of PPA-VPF. Regarding the molecular mechanism of photo-breakdown, singlet oxygen has been reported to directly oxidize methionine, tryptophan, and cysteine residues, leading to protein backbone cleavage or aggregation [22,25]. Additionally, PD-L1 contains critical residues that maintain its structural integrity [39], and oxidation of these residues by ¹O₂ may disrupt disulfide bonds, triggering irreversible denaturation.

To explore the long-term degradation mechanism, we monitored the time evolution of PD-L1 distribution after PPA-VPF incubation using confocal microscopy (Fig. 3g). At 1-h post-incubation), PD-L1 was predominantly located on the plasma membrane, while increased cellular internalization was observed 8-h later, coinciding with the internalization of PPA-VPF. This internalization may explain the slight reduction in PD-L1 abundance resulting from PPA-VPF treatment in the dark (Fig. 3c). A similar autophagic degradation mechanism has been previously demonstrated with substituted 2-amino-1-oxa-3,4-diazoles targeting the PD-1/PD-L1 pathway [40]. Moreover, the application of the lysosomal inhibitor chloroquine significantly blocked the long-term degradation of PD-L1, while having minimal effect on the early degradation (Fig. 3e). To determine whether the proteasome pathway contributes to PD-L1 degradation, we treated cells with MG132 (10 μM) during PPA-VPF photo-irradiation. The finding that MG132 did not rescue PD-L1 loss after irradiation (Fig. S4f) suggests that the proteasome pathway may not be involved in the sustained degradation of PD-L1. These results demonstrate that both immediate photo-breakdown and long-lasting autophagic degradation are present during the photodegradation of PD-L1 by PPA-VPF (Fig. 3h).

2.4. PPA-VPF induces immunogenic cell death

Since some PPA-VPF molecules can enter cells (Fig. 2a and b), we anticipated that they would generate non-proximity-generated singlet oxygen, resulting in cell death similar to VPF. Cell viability measurements showed that PPA-VPF demonstrated potent toxicity to MC38-WT cells, with an IC₅₀ of approximately 5 μM (Fig. 4a). In contrast, PD-L1-KO MC38 cells were minimally affected by 20 μM PPA-VPF under the same irradiation conditions. Moreover, PPA-VPF effectively killed PD-L1-high MDA-MB-231 cells with an IC₅₀ of approximately 1 μM, while PD-L1-low MCF-7 cells were less sensitive to PPA-VPF treatment (Fig. 4b). Notably, normal 293T cells, which do not express PD-L1, were hardly affected by 20 μM PPA-VPF under the same irradiation conditions (Fig. 4c). Conversely, VPF exhibited potent toxicity to 293T cells, with an IC₅₀ of approximately 0.2 μM. These results demonstrate the selective targeting of PPA-VPF to PD-L1-expressed cells and its enhanced safety compared to non-targeted VPF.

Fig. 4.

PPA-VPF resulting in immunogenic cell death in cancer cells. (a) Dose-dependent cytotoxicity of PPA-VPF under irradiation (5 min twice) in MC38 and PD-L1-KO-MC38 cells, and (b) in MDA-MB-231 (PD-L1 high) and MCF-7 (PD-L1 low) cells. (c) Dose-dependent cytotoxicity of PPA-VPF or VPF under irradiation (5 min twice) in normal 293T cells. HMGB1 release (d) and ATP release (e) from 4T1, MCF-7 and MDA-MB-231 cells photo-irradiated with VPF or PPA-VPF. (f-g) Maturation of BMDCs cocultured with dying 4T1 cells photoirradiated with VPF or PPA-VPF. Irradiation was performed using a 300 W Xenon arc lamp (600 nm bandpass filter, 1.5 mW/cm²) for the designated time. PPV was tested for comparison, and LPS (100 ng/mL) was used as a positive control.

Previous studies, including ours, have demonstrated that VPF-induced cell death under irradiation is immunogenic both in vitro and in vivo [21,41]. Therefore, we examined the immunogenicity of cell death caused by PPA-VPF treatment. We found that the concentrations of released ATP and high mobility group protein B1 (HMGB1)—two typical DAMPs that activate dendritic cells (DCs)—were significantly elevated in PPA-VPF-treated 4T1 cells (Fig. 4d and e). Similarly, the release of ATP and HMGB1 in both MDA-MB-231 and MCF-7 cells after irradiation was significantly enhanced compared to that in the non-treated cells. In general, PPV-VPF and VPF induced comparable levels of ATP and HMGB1 in all three cell lines, a higher release of HMGB1 from PPV-VPF was observed at a concentration of 2 μM. Therefore, we assume that the release of ATP and HMGB1 were primarily attributed to the cytotoxicity induced by VPF, whether irradiated independently or as part of the chimeric PPV-VPF. Furthermore, in vitro coculture stimulation demonstrated that dying cells from PPA-VPF treatment could trigger the maturation of murine bone marrow-derived dendritic cells (BMDCs), as evidenced by increased expression of MHC II and CD86 in CD11c⁺ DCs (Fig. 4f and g). These measurements reveal that PPA-VPF retains the ability to induce ICD, similar to VPF.

2.5. PPA-VPF exhibits potent antitumor activity in 4T1-Engrafted mice

To investigate the tumor targeting of PPA-VPF in vivo, we examined its metabolism in mice engrafted with 4T1 tumors using in vivo fluorescence imaging. Following a tail vein injection, PPA-VPF circulated systemically, with the highest fluorescence intensity observed approximately 8 h post-injection (Fig. 5a). A significant portion of the fluorescence remained detectable even after 48 h. In contrast, VPF was metabolized much more rapidly, peaking at approximately 15 min after injection, with only a small amount remaining after 120 min. The prolonged metabolism of PPA-VPF can be attributed to the proteolysis resistance of the D-peptide PPA. Additionally, we examined the distribution of PPA-VPF and VPF in the mice and found that PPA-VPF was more concentrated in tumor tissues compared to other organs, including the heart, liver, kidney, stomach, and lung, 24 h after injection (Fig. 5b). In contrast, VPF was detected in both tumor tissues and other organs, such as the liver and kidney, 60 min post-injection. These results, consistent with our in vitro observations (Fig. 2), demonstrate the effective tumor targeting of PPA-VPF in vivo.

Fig. 5.

Tumor targeting of PPA in Balb/c mice engrafted with 4T1 tumors. (a) In vivo near-infrared fluorescence imaging of mice injected with PPA-VPF or VPF at the indicated time points. (b) Distribution of VPF and PPA-VPF in mice. PPA-VPF and VPF were injected via the tail vein, and imaging was obtained based on the intrinsic fluorescence of the photosensitizer.

We then evaluated the antitumor activity of PPA-VPF in mouse models with bilateral tumors. In this study, distant tumors were induced by injecting tumor cells into the opposite side of the same mouse's back, seven days after establishing the primary tumor through subcutaneous transplantation. This setup simulates the metastatic process, allowing us to investigate the systemic immune response and the treatment's effect on metastatic sites. The light irradiation induces immunogenic cell death in the primary tumor, releasing tumor antigens that prime the immune system to recognize and attack distant metastases. 4T1 cells, which are not sensitive to ICBs [42], were engrafted in both flanks of immunity-competent BALB/c female mice. The compounds were administered via tail vein injection, but only the primary tumor in each mouse was irradiated with a 689 nm laser (100 mW/cm² for 8 min) at either 24 h (PPA-VPF group) or 0.5 h (VPF group) post-injection (Fig. 6a). Treatments were repeated three times every two days.

Fig. 6.

Antitumor activity of PPA-VPF in mice engrafted with bilateral 4T1 tumors. (a) Procedures of tumor treatment. Growth curves, tumor growth inhibition rate and images of dissected xenografts of primary tumors (b) and distant tumodrs (c) with the indicated treatment in Balb/c mice xenografted with 4T1. (d) H&E staining and TUNEL analysis of the primary tumors after the indicated treatment. (e) Growth curve of 4T1 tumors in Balb/c nude mice with the indicated treatment. (f) Frequency of CD8⁺CD3⁺ T cells in tumor-infiltrating lymphocytes and frequency of IFN-γ⁺CD8⁺ effector T cells in total CD8⁺ T cells collected from mice tumors after the indicated treatment (Fig. S6). Quantification of relative PD-L1 content based on western blotting (g) and immunofluorescent staining of PD-L1 (h) in the primary tumors at the endpoint of the indicated treatment.

As expected, the peptide PPA had a minimal effect on the growth of 4T1 tumors at a dose of 2 mg/kg (Fig. 6b and c, S5a). VPF could mildly inhibit the growth of both primary and distant tumors by triggering antitumor immunity, consistent with previous reports [41]. However, PPA-VPF treatment not only significantly inhibited the growth of primary tumors but also distant tumors (Fig. 6b and c). The tumor growth inhibition (TGI) rates of PPA-VPF for primary and distant tumors were approximately 74% and 48%, respectively, which are higher than the corresponding rates for VPF treatment (39% and 35%, respectively). We also used pathological and immunofluorescence staining to analyze the primary tumors after the indicated treatments. Hematoxylin and eosin (H&E) staining revealed that tumor tissues treated with PPA-VPF showed significantly weakened nucleus-to-cytoplasm interactions (Fig. 6d). The TUNEL assay confirmed that more tumor cells underwent apoptotic death in PPA-VPF-treated tumors than in VPF-treated tumors. In contrast, no therapeutic effects of PPA-VPF on the distant tumor were observed in the immune-deficient Balb/c nude mice, while its inhibitory effect on the primary tumor was evident due to the direct PDT cytotoxicity (Fig. 6e). These results point to the essential role of immune activity for PPA-VPF to exert its inhibitory effect on distant tumors.

To assess this antitumor immune response in vivo, we measured the abundance of PD-L1 in primary 4T1 tumor tissues at the treatment endpoint using both western blotting and immunofluorescent staining. Remarkably, treatment with PPA-VPF in vivo led to a significantly greater reduction in PD-L1 expression compared to treatments with PPA or VPF alone (Fig. 6g and h, S5c), consistent with our in vitro findings (Fig. 3).

Furthermore, we directly assessed the lymphocyte activity induced by PPA-VPF in 4T1-bearing Balb/c mice, following treatment procedures similar to those for the bilateral model (Fig. S6a). Three days after the final treatment, we collected tumor tissues for immunological evaluation using flow cytometry. PPA-VPF treatment resulted in a frequency of cytotoxic T cells (CTLs, CD8⁺CD3⁺) in tumor-infiltrating lymphocytes as high as ∼55%, significantly higher than the 40% observed with VPF treatment and ∼20% with PPA treatment (Fig. 6f–S6b). Additionally, the functional activity of CD8⁺ T cells, as indicated by IFN-γ expression, increased to ∼55% in the PPA-VPF group, compared to ∼33% in the VPF group and ∼25% in the PPA group (Fig. 6f–S6c). Overall, PPA-VPF demonstrated the highest efficacy in triggering antitumor immunity, although both VPF and PPA also induced enhanced immune activation compared to PBS. These differences in immunological evaluation correlated well with their antitumor efficacy in the bilateral 4T1 model (Fig. 6b and c).

The clinically approved drug VPF exhibits good safety, partially due to its rapid metabolism [43]. We were interested in the biosafety of PPA-VPF, whose metabolism is significantly prolonged due to the presence of PPA. The body weights of mice (Fig. S5c) in all treated groups did not show significant changes during treatment. H&E staining of the main tissues (heart, liver, spleen, lung, and kidney) from treated mice revealed that PPA-VPF caused no obvious damage to these organs, while renal damage was observed in VPF-treated mice (Fig. S7a). Moreover, we performed serum biochemistry assays on healthy Balb/c mice after intravenous administration of compounds (4 mg/kg PPA, 2 mg/kg VPF, and 6 mg/kg PPA-VPF) three times every two days (without light irradiation) (Fig. S7b–S7i). Compared with the control group, the PPA-VPF-treated group and the VPF-treated group exhibited similar levels of biochemical indices, including alanine aminotransferase (ALT), aspartate aminotransferase (AST), albumin (ALB), total protein (TP), creatinine (CR), urea (UREA), and lactate dehydrogenase (LDH). These results indicate that all treatments did not affect the functions of the liver and kidneys in the mice. Therefore, PPA-VPF demonstrated excellent biocompatibility and biosafety for antitumor therapy.

2.6. PPA-VPF exhibits potent antitumor activity in MC38-Engrafted and CT26-Engrafted mice

We investigated the antitumor efficacy of PPA-VPF in mice engrafted with MC38 and CT26 tumors. In mice with MC38-WT tumors, PPA-VPF monotherapy significantly reduced tumor growth compared to PBS controls (Fig. 7a and b). However, in MC38-PD-L1-KO mice, which exhibited slower tumor growth than those with MC38-WT tumors, PPA-VPF did not show a significant effect. These findings underscore the crucial role of PD-L1 targeting in the effectiveness of PPA-VPF.

Fig. 7.

Therapeutic response in MC38 and CT26 xenograft models. (a) Tumor growth curves from C57BL/6J mice (n = 4) with primary tumor samples. Treatment groups: vehicle control (PBS), PD-L1 KO, PPA-VPF (8 mg/kg), and PD-L1 KO with PPA-VPF. Light irradiation was performed using a 689 nm laser 24 h (PPA-VPF) or 30 min (VPF) after injection (100 mW/cm² for 8 min). (b) The xenografts obtained at the endpoint of the indicated treatments. (c) Tumor growth curves with representative primary tumor specimens from BALB/c mice (n = 4). (d) The images of dissected xenografts of primary tumors and distant tumors with the indicated treatment. Treatment groups: vehicle control (PBS), monotherapy PD-L1 antibody (5 mg/kg every three days over a 12-day course), monotherapy PPA-VPF (8 mg/kg every two days, administered thrice), combination therapy (PPA-VPF + PD-L1 antibody). Data represent mean ± SD. (e) IHC staining and analysis of CD3 expression in CT26 primary tumor following treatment PPA-VPF and PD-L1 antibody. Scale bar represents 10 μm.

We further assessed the therapeutic effects of PPA-VPF in combination with an anti-PD-L1 monoclonal antibody (mAb) using the PD-1/PD-L1 blockade-refractory CT26 bilateral tumor model. Monotherapy with either PPA-VPF or the PD-L1 mAb demonstrated limited efficacy in inhibiting the growth of primary tumors (Fig. 7c and d). However, the combination of PPA-VPF and the PD-L1 mAb exhibited remarkable synergistic activity, resulting in significant growth inhibition. Notably, both PPA-VPF alone and its combination with the PD-L1 mAb significantly inhibited the growth of distant tumors (Fig. 7d). This synergy highlights the potential of combining PPA-VPF with PD-1/PD-L1 blockade to overcome resistance to immune checkpoint inhibitors.

To clarify the absolute number of intratumoral T cells, we conducted IHC staining for CD3, a pan-T cell marker, in these tumor tissues. CT26 tumors displayed a negligible number of CD3⁺ T cells. Both PPA-VPF and the PD-L1 mAb alone significantly increased CD3⁺ T cell density and an additional increase was observed when they were combined (Fig. 7e), consistent with the trends observed for ICB treatment [44].

These findings further emphasize the dual-action mechanism of PPA-VPF treatment in immunotherapy. On one hand, the ICD effects of PPA-VPF (Fig. 4d–g) lead to DC activation and T cell recruitment. On the other hand, it acts as an immune checkpoint inhibitor to effectively degrade tumor PD-L1 (Fig. 6g and h) and maintain T cell activity (Fig. 6f and 7e). Additionally, the synergistic effect of PPA-VPF and the PD-L1 mAb highlights the potential of combining PD-L1-targeted photodegradation with current immunotherapy to counteract tumor-induced immunosuppression.

3. Conclusions

We have demonstrated a novel PDTAC strategy that utilizes a dual mechanism to trigger potent antitumor immunity. By conjugating a PD-L1-targeted D-peptide with PPA-VPF, we enhanced the tumor targeting of VPF. The D-type targeting peptide's resistance to proteolysis allows PPA-VPF to exhibit improved metabolic stability compared to the parent VPF. Notably, PPA-VPF effectively degrades PD-L1 both in vitro and in vivo. The photodegradation of PD-L1 occurs through two pathways: immediate protein photolysis and long-lasting lysosomal-dependent degradation. Furthermore, PPA-VPF induces cancer cell death and results in ICD, stimulating the maturation of DCs. Following intravenous administration, PPA-VPF demonstrated potent antitumor efficacy against both primary and distant tumors in several tumor-bearing mouse models. PPA-VPF treatment activated robust adaptive immunity, as indicated by an increase in CD8⁺ T cells and mature DCs in vivo, following the degradation of PD-L1 and the induction of ICD. In summary, our PD-L1-targeted chimera may offer insights for developing a novel strategy to enhance response rates to ICB in cancer immunotherapy.

4. Experimental section

General. All chemicals and solvents were commercially purchased and used without further purification. All solvents were reagent grade or HPLC grade. The purity of all synthesized peptides and PDTACs was >95% based on analytical HPLC.

Chemicals. Fmoc-D-Asn(Trt)-OH, Fmoc-D-Tyr(OtBu)-OH, Fmoc-D-Ser(tBu)-OH, Fmoc-D-Lys(Boc)-OH, Fmoc-D-Pro-OH, Fmoc-D-Thr(tBu)-OH, Fmoc-D-Asp(OtBu)-OH, Fmoc-D-Arg(Pbf)-OH, Fmoc-D-Gln(Trt)-OH, Fmoc-D-His(Trt)-OH, Fmoc-D-Phe-OH, HOBt and HATU were purchased from GL Biochem. Diisopropylethylamine (DIEA) and trifluoroacetic acid (TFA) were purchased from J&K. Piperidine was purchased from Energy Chemical. 9-[(9H-Fluoren-9-ylmethoxy) carbonylamino]- 4,7-dioxanonanoic acid (Fmoc–NH–PEG₂- CH₂CH₂–COOH), 1-(9H-Fluoren-9-yl)-3-oxo-2,7,10,13,16,19-hexaoxa-4-azadocosan-22-oic acid (Fmoc–NH–PEG₅- CH₂CH₂–COOH), Fmoc–NH–PEG₁₀-acid (Fmoc–NH–PEG₅- CH₂CH₂–COOH) and 6-{[(9H-Fluoren-9-ylmethoxy)carbonyl]amino}.hexanoic acid (Fmoc-ℇ-Acp-OH) were purchased from Bidepharmatech. Verteporfin was purchased from Macklin. 4-Hydroxy-2,2,6,6-tetramethylpiperidine (4-OH-TEMP) and trifluoroethanol (TFE) were purchased from Sigma. DAPI was purchased from Bestbio.

Synthesis and purification of PDTACs. The targeting peptide (^DN^DY^DS^DK^DP^DT^DD^DR^DQ^DY^DH^DF) in this study was synthesized using Fmoc (9-fluorenylmethyloxycarbonyl)-solid-phase peptide chemistry with Rink-Amide resin (Shimadzu). A coupling system using 2-(7-azabenzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium hexafluorophosphate (HATU)/1-hydroxybenzotriazole (HOBt)/N,N-diisopropylethylamine (DIEA) was employed. Fmoc–NH–PEG_n-CH₂CH₂–COOH (n = 2, 5, 10) or Fmoc-ℇ-Acp-OH as a linker unit was linked to the N-terminus of the peptide chains. After Fmoc protecting group removal, the photosensitizer verteporfin ((±)-trans-3,4-dicarboxy-4,4a-dihydro-4a,8,14,19-tetramethyl-18-vinyl-23H,25H-benzo[b]porphine-9,13-dipropanoic acid) was further coupled to the N-terminus of the linker using HATU/1-hydroxy-7-azabenzotriazole (HOAT)/DIEA. After synthesis, deprotection of the protecting groups and detachment of peptide-photosensitizer conjugations from the resin were performed using trifluoroacetic acid (TFA)/triisopropylsilane (TIPS)/H2O (95/2.5/2.5) at 20 °C for 2 h followed by purification via reverse-phase high-performance liquid chromatography (RP-HPLC). The whole experimental process was protected against exposure to light.

All HPLC separations involved a mobile phase of 0.05% (v/v) TFA in water (solvent A) and 0.04% (v/v) TFA in acetonitrile (solvent B). Analytical LC‒MS analyses were performed using a Water Alliance e2695 Separations Module equipped with an Agilent C18 column (5.0 μm, 4.6 × 150 mm, 0.4 mL/min), a Water 2489 UV/Visible Detector and a Waters SQD mass spectrometer (Alliance e2695-SQD). The wavelengths of the UV detector were set to 210 nm and 220 nm. Preparative HPLC separations were performed using two Shimadzu LC-20AR semipreparative solvent delivery units, a Shimadzu SPD-20A UV detector and a Shimadzu CBM-20A system controller equipped with a Dr. Maisch ReproSil 300 C18 column (5.0 μm, 20 × 250 mm) at a flow rate of 12 mL/min. The wavelengths of the UV detector were set to 210 and 220 nm.

The photodegradation-targeting chimeras exhibited two positional isomers derived from VPF, which had been observed in LC‒MS (Fig. S1, Fig. S8-S11). The two isomers were mixed at 1:1 for all further analyses.

Analysis of morphology, size distribution and zeta potential. The morphologies of the chimeras were observed on a transmission electron microscope (JEM-1400 PLUS). Chimeras were prepared in PBS buffer (pH 7.4). Briefly, 8 μL of prepared chimera solutions was loaded onto the copper grids of carbon support films (200 meshes) for 1 min. Excess gel was removed by blotting with filter paper, followed by the addition of 8 μL of negative stain (2% aqueous phosphotungstic acid) and staining for 1 min. After removing the remaining liquid with filter paper, copper grids were allowed to dry in air overnight prior to observation. The size distribution of the chimeras in PBS buffer (pH 7.4) was measured on a Nano-ZS90 Zetasizer by dynamic light scattering (DLS, Malvern Instruments). The zeta potentials were simultaneously measured in PBS.

Absorption and fluorescence spectrometry. The UV-Vis absorption and fluorescence spectra (λ_ex: 532 nm) of VPF or PPA-VPF were measured with a UV-Visible spectrophotometer (Yok Instruments) and fluorescence spectrometer (Cary Eclipse), respectively.

Measurement of singlet oxygen by EPR. Singlet oxygen generated by PPA-VPF or VPF was trapped by the spin scavenger 4-hydroxy-2,2,6,6-tetramethylpiperidine (4-OH-TEMP, 200 mM) during photoexcitation and was then detected on an X-band Bruker ER A200 spectrometer. Briefly, approximately 500 μL of sample in a 1.5 ml centrifuge tube was irradiated by a 300 W Xenon arc lamp (CEAULIGHT) with a 600 nm bandpass filter (∼1.5 mW/cm²) for the designated time. To detect singlet oxygen, a 200 mM final concentration of 4-OH-TEMP was added to the PPA-VPF or VPF solution (20 μM) before irradiation. After irradiation, a 30 μL aliquot was immediately aspirated into a glass capillary and transferred to the EPR resonator. The mixture solvent was PBS or 50% ACN (PBS: acetonitrile = 1:1). Typical settings applied for EPR detection were as follows: scan range, 100 G; sweep time, 60 s; modulation amplitude, 1 G; modulation frequency, 100 kHz; and microwave power, 19.23 mW.

Surface plasmon resonance (SPR) measurement. The affinities between hPD-L1 and chimeras were measured by SPR (Biacore 8K, GE Healthcare, Sweden), all at 25 °C. The purified active hPD-L1 was diluted in 10 mM sodium acetate buffer (pH 4.5, GE Healthcare), resulting in a protein concentration of 40 μg/mL. The diluted hPD-L1 was soon covalently immobilized to flow cell 2 of a new sensor chip (Series S Sensor Chip CM5, GE Healthcare) via the primary amine group using the standard Amine Coupling Kit (GE Healthcare). The target hPD-L1 immobilization level was 8000 RU. Flow cell 1 was blank as a reference, without any modification. Measurements were run at a flow rate of 30 μL/min in PBS-P buffer (10 mM phosphate buffer with 2.7 mM KCl, 137 mM NaCl, and 0.05% surfactant P20, final pH 7.4, GE Healthcare) or HEBS buffer containing 0.05% surfactant tween 20, which was vacuum-filtered and degassed immediately before the experiment. To obtain the data for kinetic and affinity analysis, a concentration gradient of chimeras was freshly prepared in PBS-P running buffer with at least five concentrations (2-fold serial dilutions), and the concentration range was optimized according to different analytes. RU values were collected, and all the experimental data were globally analyzed by a steady-state model within Biacore 8K Evaluation Software, version 1.0.

Coomassie Brilliant Blue staining. The hPD-L1 protein (100 μg/mL) was treated with PPA-VPF (10 μM) or VPF (10 μM) at 25 °C and then irradiated for different durations (1.5 mW/cm2). The treated solution was separated by SDS-PAGE in 12.5% polyacrylamide gels and then stained with Coomassie brilliant blue. To prepare a solution of 0.025% Coomassie R-250, 1 Phast Gel Blue tablet was dissolved in 1.6 L of Coomassie R-250 in 1 L of 10% (v/v) acetic acid. The solution was heated to 90 °C and poured over the gel in a stainless-steel tray, and then, the tray was covered with a lid on a laboratory shaker for 10 min. The gel was destained using 10% (v/v) acetic acid for at least 2 h at room temperature.

Cell Culture. MDA-MB-231, MCF-7 and 4T1 were purchased from the National Infrastructure of Cell Line Resource of China and tested negative for mycoplasma contamination. CT26, MC38 colon adenocarcinoma and CD274 Knockout cell line (MC38) cell lines were purchased from the Ubigene (Guangzhou, China). MCF-7, 4T1 and MC38 cells were cultured in Dulbecco's modified Eagle medium (DMEM) with 10% FBS, 100 U ml⁻¹ penicillin, and 100 μg ml⁻¹ streptomycin (all from M&C GENE TECHNOLOGY). CT26 and MDA-MB-231 cells were cultured in Roswell Park Memorial Institute (RPMI) 1640 with 10% Certified Fetal Bovine Serum, FBS (VivaCell, Shanghai, China), 100 U ml⁻¹ penicillin, and 100 μg ml⁻¹ streptomycin (all from M&C GENE TECHNOLOGY). These cells were maintained at 37 °C in a humidified 5% CO₂ incubator (ESCO), and subculture was conducted every 2-3 days. Cell freezing medium was purchased from Shanghai Chuanqiu. Biotechnology Co., Ltd., China.

Assay of cellular internalization of photosensitizers. MDA-MB-231 cells were plated on 24-well plates and treated with VPF or PPA-VPF at different concentrations for 8 h at 37 °C with or without red-light irradiation. After incubation, the cells were harvested and resuspended in 1 × PBS for flow cytometry analysis (Beckman CytoFlex) (λ_ex: 638 nm). The change in red fluorescence intensity was quantified by FlowJo. The results are expressed as the mean fluorescence ± SD (n = 3).

To investigate the endocytosis pathways, the inhibitor CPZ (20 μM), MβCD (5 mM), or colchicine (100 μM) (Targetmol, USA.) was first coincubated with cells for 1 h. Then, the inhibitors were replaced by PPA-VPF and further coincubated for 8 h. The harvested cells were washed twice with PBS and resuspended in 500 μL of PBS. Internalized PPA-VPF was measured using flow cytometry (BD FACSCalibur) (λex: 631 nm). A total of 10,000 cells in each sample were analyzed.

Immunofluorescence analysis of PD-L1 expression in cells. MDA-MB-231 cells or MCF-7 cells were seeded on a 24-well glass plate at the bottom of wells covered with round coverslips at a density of 5 × 10⁴ cells per well. After the cells were attached to the surface of the round coverslip, the solution containing PPA-VPF or VPF was added to incubate the cells for 1 h and then incubated for different times after red-light irradiation. Then, the medium in the wells was removed. Cells attached to the coverslips were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 10 min at room temperature. The solution in the wells was removed, and the cells were washed three times with PBS buffer for 1 min each time. BSA (5%) in PBS was used as a blocking solution to incubate the cells for 1 h to reduce interference from the fluorescent background. The blocking solution was aspirated, and the washing step mentioned above was repeated three times. Alexa Fluor 488-conjugated PD-L1 antibody diluted 25 times in 5% BSA in PBS buffer was used to incubate the cells for 1.5 h at room temperature. The primary antibody solution was removed, and the cells were washed three times with PBS buffer and then stained with DAPI (0.5 μg mL−1, Beijing Solarbio Science & Technology Co., Ltd.) for 4 min at room temperature. All operations were protected from light. All images were taken using a laser scanning confocal microscope at the same voltage.

Analysis of PD-L1 degradation by flow cytometry. Cells were seeded into 24-well plates and allowed to reach 60% confluence. Then, the MDA-MB-231 cells were treated with DMSO, PPA, PPA-VPF, VPF or PPA-VPF + chloroquine (Molnova Chemicals, Shanghai China) (2 μM, chloroquine 10 nM) at 37 °C for 1 h and then irradiated with a 300 W Xenon arc lamp (600 nm bandpass filter, 1.5 mW/cm², 6 min) or without irradiation with the following incubation for different times. We removed the cells into different centrifuge tubes with the trypsin digestion method. After incubation with PE-conjugated anti-human PD-L1 antibody (BioLegend) for 30 min at room temperature, the cells were centrifuged (800×g, 3 min), and the supernatant was decanted. Cells were resuspended in 1 mL of cold staining buffer by pipetting gently for 5 min and centrifuged at 800×g for 3 min at 4 °C. Then, the cells were washed with cold staining buffer three times. Finally, the stained cells were analyzed by FACS (BD FACSCalibur) (λex: 488 nm) after resuspension with 300 μL staining buffer. The whole process was protected from exposure to light.

Analysis of PD-L1 expression using western blotting. The expression level of PD-L1 in MDA-MB-231 cells was tested using western blotting assays with PD-L1 polyclonal antibodies from three different brand (Abcam, Proteintech and Cell Signaling Technology). MDA-MB-231 cells were seeded on cell culture dishes with a diameter of 10 mm and cultured for 24 h. After the cells grew to a density of ≈80%, the cells were treated with DMSO, PPA (1 μM), PPA-VPF (1 μM or 3 μM) or VPF (1 μM) (5 min x 2) at 37 °C for 1 h and then irradiated with a 300 W Xenon arc lamp (600 nm bandpass filter, 1.5 mW/cm²) with incubation for 8 h. Then, we removed the drug-containing medium and washed the cells three times with PBS buffer. The cells were harvested and lysed in cell lysis buffer containing protease inhibitors, which were provided by a cell membrane protein extraction kit. After quantitative analysis using bicinchoninic acid (BCA), equal amounts (30 μg) of the samples were separated using sodium dodecyl sulfate‒polyacrylamide gel electrophoresis (SDS‒PAGE) and then transferred to a polyvinylidene difluoride (PVDF) membrane. The membrane was blocked with 5% skim milk to eliminate any nonspecific interference. The membranes were incubated with PD-L1 polyclonal antibody and GAPDH antibody at 4 °C overnight followed by incubation with HRP-labeled secondary antibody for 1.5 h. The membranes were finally detected using Immobilon Western HRP substrate and Chemiluminescent Imaging System.

PDT treatment and cell viability assay. Cells were seeded onto 96-well plates (3000 cells per well) and allowed to adhere overnight. The medium was replaced on the following day with 100 μL of growth medium containing a required concentration of VPF or PPA-VPF, and the cells were incubated in the dark. One hour later, the cells were exposed to red light irradiation using a 300 W Xenon arc lamp (600 nm bandpass filter, 1.5 mW/cm²) for 5 min twice and were further incubated thereafter. Cell viability was typically assessed 24 h after light irradiation using a CCK-8 kit (Life-iLab, Shanghai, China). All cell viability data were normalized to the DMSO vehicle condition. Measurements were performed with three independent replicates.

Animals. All animal experiments were performed according to the protocols stipulated by the Guideline for Care and Use of Laboratory Animals of Peking University and approved by the Animal Ethics Committee of Peking University (LA2021274). Female BALB/c mice (6–8 weeks old) and female BALB/c nude mice (6-8 weeks old) were purchased from the Department of Animal Science of Peking University Health Science Center and housed in specific pathogen-free conditions.

In vivo tumor distribution of PPA-VPF or VPF. 4T1 cells (5 × 10⁵) were subcutaneously injected into the right flanks of BALB/c mice. As the tumor volume reached approximately 200 mm³, mice were intravenously administered PPA-VPF (6 mg/kg) or VPF (2 mg/kg). An in vivo near-infrared fluorescence imaging system was used for PPA-VPF or VPF distribution detection at different time points after injection with an IVIS imaging system coupled to IVIS spectrum software (CALIPER).

In vivo antitumor studies. A bilateral tumor model was established for in vivo antitumor studies. 4T1 cells (5 × 10⁵) were subcutaneously injected into the right flanks of female BALB/c or male C57BL/6J mice to create primary tumors; 1 × 10⁵ cells were subcutaneously injected into the left flanks to create distant tumors. As the primary tumor volume reached ∼100 mm³, the mice were randomly divided into four groups (4 in each group, 16 in total) using a computer based random order generator, and intravenously injected with PBS, PPA (4 mg/kg), PPA-VPF (6 mg/kg), and VPF (2 mg/kg). The mice in the third and fourth groups were selectively irradiated on primary tumors by a 689 nm laser 24 h or 30 min after injection (100 mW/cm², 8 min). The treatments were repeated every 2 days three times. The weights and tumor volumes were recorded. Additionally, serum biochemistry assays were conducted to evaluate the influence of PPA-VPF or VPF on heart, liver and kidney functions. After the treatments were finished, sera were harvested to detect the liver- and kidney-related biochemical indexes. Finally, mice were euthanized on day 11 for pathologic and immunofluorescence evaluation. In addition, the heart, liver, spleen, lung, and kidney were collected for further histopathological analysis.

The same experiments were also repeated in female BALB/c nude mice (7-8 weeks old) to examine the involvement of immune system in the antitumor effects.

In vivo immunological evaluation of PPA-VPF. 4T1 cells (5 × 10⁵) were subcutaneously injected into the right flanks of female BALB/c mice. As the tumor volume reached approximately 300 mm³, the mice were randomly divided into four groups (3 in each group, 12 in total) using a computer based random order generator. The groups are PBS, PPA (4 mg/kg), PPA-VPF (6 mg/kg), and VPF (2 mg/kg). Mice in the third and fourth groups were selectively irradiated by a 689 nm laser 24 and 0.5 h after injection for 8 min at an intensity of 100 mW/cm². Each treatment was performed in triplicate. On the third day after the final treatment, the mice were euthanized, and the spleen and tumor tissues were dissected for immunological evaluation. To investigate the DC maturation frequency, DCs were harvested from the spleen and stained with anti-CD11c APC (BioLegend), anti-CD86 APC-Cy7 (BioLegend), and anti-CD80-PE (BioLegend) antibodies. The frequency of CD11c⁺/CD80⁺/CD86⁺ DCs was detected by flow cytometry. For the tumor-infiltrating lymphocyte investigation, mice were sacrificed, and tumors were harvested. Tumor tissue was isolated and added to a solution of collagenase (400 units/mL), DNase 1 (100 μg/mL), and hyaluronidase (0.04 unit/mL). The mixture was shaken at 37 °C for 1 h. Then, the mixture was passed through a 70 μm nylon cell strainer. The sample was washed three times with PBS (with 2% FBS) and then resuspended at a density of 1 × 10⁶ cells/mL for flow cytometry analysis. The collected lymphocyte suspensions were blocked with anti-CD3-FITC and anti-CD8-PE-Cy7 antibodies at 4 °C for 30 min for flow cytometry analysis. For IFN-γ⁺/CD8⁺ T-cell analysis, the harvested lymphocyte suspensions were blocked with fluorescence-labeled anti-CD3-FITC and anti-CD8-PE-Cy7 antibodies, permeabilized in Perm/Wash buffer (BD Biosciences, USA) for 30 min and stained with anti–IFN–γ-PE antibodies. For quantification, the proportion of each immune subpopulation was determined by flow cytometry.

TUNEL cell apoptosis staining. Paraffin sections of tumor tissues were deparaffinized in a gradient. After the slices were slightly dried, the working solution of proteinase K was added to cover the tissues, and the slices were incubated at 37 °C for 22 min and washed three times with PBS buffer for 5 min each. Then, the working solution of membrane breaking was dropped to cover the tissues, which were incubated for 20 min at room temperature, and the above cleaning step was repeated. Following the protocol provided by the kit, the reaction solution was added to cover the tissues, and the slices were incubated at 37 °C for 2 h. After washing three times with PBS buffer, DAPI staining solution (Cat No. 40728ES03, Yeasen, Shanghai, China) was dropped on the slices and incubated for 10 min at room temperature in the dark. Finally, after repeating the above cleaning steps, the slides were mounted with antifluorescence quenching mounters. The obtained slices were observed under a fluorescence microscope, and images were collected.

Immunofluorescence staining of tumors. Isolated tumors were fixed in 4% paraformaldehyde for 24 h at room temperature, dehydrated, and embedded in paraffin wax. Before being stained with an antibody, tissues were transferred to slides and deparaffinized. The anti-mouse PD-L1 antibody (Cell Signaling Technology) was added to the slides and incubated at 4 °C overnight for immunofluorescence staining. After being washed three times with PBS, tissues were incubated with DAPI. Immunofluorescence images were acquired on a confocal microscope.

Immunohistochemical Staining of tumors. Formalin-fixed paraffin-embedded tumor sections (4 μm) were deparaffinized, rehydrated, and subjected to antigen retrieval in citrate buffer (pH 6.0, 95 °C, 20 min). Endogenous peroxidase was blocked with 3% H₂O₂, followed by 5% goat serum blocking. Sections were incubated with anti-CD3 (1:200, Selleck) overnight at 4 °C, then treated with HRP-polymer system (Dako) and developed with DAB. Nuclei were stained with hematoxylin. Negative controls omitted primary antibody. Images were captured under a light microscope, and CD8⁺ T cells were quantified via ImageJ (5 random fields/section).

Measurement of released ATP and HMGB1. 4T1 cells were treated with PPA-VPF or VPF in medium with 2% FBS. Then, the supernatants were collected and centrifuged at 15,000 rpm at 4 °C for 3 min. The supernatants were either stored at − 80 °C or used immediately for ATP measurements. ATP analysis was performed using the CellTiter-Glo® Luminescent Cell Viability Assay kit (Promega, G7571) as described by the manufacturer. The luminescence was measured on a BioTek Synergy Neo2.

HMGB1 was measured by an ELISA kit (Elabscience Biotechnology Co., Ltd.). After the indicated time points, the supernatant was collected and cleared from dying tumor cells by centrifugation and frozen at −20 °C. All assays were performed in accordance with the respective manufacturers’ instructions, and HMGB1 was quantified using BioTek Synergy Neo2. The data were analyzed with a four-parameter logistic curve fit.

Analysis of BMDC maturation. Over 7 days, bone marrow-derived dendritic cells (BMDCs) were differentiated from the femurs and tibias of C57BL/6 J mice at the age of 7-9 weeks using RPMI medium (GIBCO) supplemented with 5% heat-inactivated fetal calf serum, 20 ng/ml mGM-CSF, 10 ng/ml IL-4, 1% l-glutamine, 50 μM 2-mercaptoethanol, and 1 mM pyruvate. Fresh culture medium was added on day 2, and on day 4, the medium was refreshed.

The obtained BMDCs were coincubated with dying 4T1 cells for 18 h. As a quality control, BMDCs were stimulated in parallel with 100 ng/ml E. coli lipopolysaccharide (LPS). The cells were then collected, centrifuged (400×g, 6 min, 4 °C), and washed once in phosphate buffered saline (PBS, Life Technologies). Maturation of BMDCs was analyzed by immunostaining with anti-CD11c APC (Biolegend), anti-CD86 APC-Cy7 (Biolegend), anti-MHCⅡ PE-Cy7 (Biolegend), and mouse Fc-block (Biolegend). True uptake of dead cells by BMDCs was determined using a gating strategy that allows analysis of only single cells.

Serum biochemistry assays. A total of 100 μL of PBS, PPA, PPA-VPF or VPF was intravenously injected into Balb/c mice. Twelve mice were randomly selected for blood collection, with three mice in each group. Orbital blood was collected to detect the liver- and kidney-related biochemical indexes after the mice were intravenously injected with PBS, PPA, PPA-VPF or VPF every 2 days for three times. The collected 500 μL blood specimens were centrifuged at 3000 r/min for 8 min to collect the supernatant for the serum test. The serum tests included measurements of alanine aminotransferase (ALT), aspartate aminotransferase (AST), albumin (ALB), total protein (TP), creatinine (CR), urea (UREA), lactate dehydrogenase (LDH), and glucose (GLU), which were all performed by relevant assay kits.

Statistical analysis. All data are represented as the mean ± SEM. All statistical analyses were calculated using GraphPad Prism 9.3.0 software and the significance was analyzed by Student's t-test or two-way ANOVA. ∗P < 0.05; ∗∗P < 0.01; ∗∗∗P < 0.001; ∗∗∗∗P < 0.0001; ns, not significant.

Notes

A patent (ZL202310952308.5) has been granted to G.L. and S.L. in China. The other authors declare no competing financial interests.

CRediT authorship contribution statement

Sijin Liu: Data curation, Investigation, Writing – original draft. Zhaoting Yang: Data curation, Investigation, Writing – review & editing. Biao Wang: Data curation, Investigation. Shuyu Huan: Data curation, Investigation. Zixi Li: Data curation, Investigation. Xunbin Wei: Supervision, Writing – review & editing. Guoquan Liu: Conceptualization, Funding acquisition, Project administration, Supervision, Writing – review & editing.

Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:

A patent (ZL202310952308.5) has been granted to G.L. and S.L. in China. The other authors declare no competing financial interests.

Appendix A. Supplementary data

The following is the Supplementary data to this article.

Data availability

Data will be made available on request.