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Prodrug-based combinational nanomedicine remodels lipid metabolism for reinforced ferroptosis and immune activation
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Ling Lina, Zaixiang Fanga, Guohao Liua, Yiwei Liua, Zhiqian Lia, Dayi Pana, Yunkun Lia, Hemi Kangb, Xiaoding Shena, Jingyao Zhanga, Qiyong Gonga, c, d, *, Kui Luoa, c, *, Jing Jinga, *
Acta Pharmaceutica Sinica B | 2025, 15(5) : 2746 - 2763
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Acta Pharmaceutica Sinica B | 2025, 15(5): 2746-2763
• ORIGINAL ARTICLES •
Prodrug-based combinational nanomedicine remodels lipid metabolism for reinforced ferroptosis and immune activation
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Ling Lina, Zaixiang Fanga, Guohao Liua, Yiwei Liua, Zhiqian Lia, Dayi Pana, Yunkun Lia, Hemi Kangb, Xiaoding Shena, Jingyao Zhanga, Qiyong Gonga, c, d, *, Kui Luoa, c, *, Jing Jinga, *
Affiliations
  • aDepartment of Radiology, Huaxi MR Research Center (HMRRC), Institution of Radiology and Medical Imaging, Breast Center, Institute of Breast Health Medicine, State Key Laboratory of Biotherapy, Core Facilities, West China Hospital, Sichuan University, Chengdu 610041, China
  • bCollege of Chemical Engineering, Sichuan University, Chengdu 610041, China
  • cFunctional and Molecular Imaging Key Laboratory of Sichuan Province, NHC Key Laboratory of Transplant Engineering and Immunology, Research Unit of Psychoradiology, Chinese Academy of Medical Sciences, Chengdu 610041, China
  • dXiamen Key Lab of Psychoradiology and Neuromodulation, Department of Radiology, West China Xiamen Hospital of Sichuan University, Xiamen 361021, China
About Author:

E-mail addresses: (Qiyong Gong),

(Kui Luo),

(Jing Jing).

†

These authors made equal contributions to this work.

Author contributions

Ling Lin, Zaixiang Fang, Qiyong Gong, Kui Luo, and Jing Jing designed the research. Ling Lin and Zaixiang Fang carried out the experiments and performed data analysis. Yiwei Liu, Guohao Liu, Zhiqian Li, Xiaoding Shen, Hemi Kang, and Jingyao Zhang participated in some of the experiments. Zhiqian Li provided experimental drugs and quality control. Ling Lin, Zaixiang Fang, Dayi Pan, and Yunkun Li wrote the manuscript. Qiyong Gong, Kui Luo, and Jing Jing revised the manuscript. All of the authors have read and approved the final manuscript.

doi: 10.1016/j.apsb.2025.03.016
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Ferroptosis is a form of programmed cell death characterized by overwhelmed lipid oxidation, and it has emerged as a promising strategy for cancer therapy. Enhanced ferroptosis could overcome the limitations of conventional therapeutic modalities, particularly in difficult-to-treat tumors. In this study, we developed a dual-modality therapy in nanomedicine by combining paclitaxel (PTX) chemotherapy and pyropheophorbide-a (Ppa) phototherapy. Heparin (HP) was grafted with poly(N-(2′-hydroxy) propyl methacrylamide) (pHPMA) using reversible addition–fragmentation chain transfer polymerization to form HP-pHPMA (HH), which was utilized to deliver Ppa and PTX, yielding HP-pHPMA-Ppa (HH-Ppa) and HP-pHPMA-PTX (HH-PTX), respectively. The prodrug-based combinational nanomedicine (HH-PP) was formed by co-assembly of HH-PTX and HH-Ppa. It was found that HH-PP treatment significantly disrupted lipid metabolism in triple-negative breast cancer (TNBC) cells, induced extensive lipid oxidation, and promoted ferroptosis. In vivo, HH-PP intervention achieved a tumor growth inhibition rate of 86.63% and activated adaptive immunity with an elevated CD8+ cytotoxic T cell infiltration level. This combinational nanomedicine offers a promising platform for co-delivery of multiple therapeutic agents. It exerts a promising anti-tumor effect via enhanced ferroptosis and ferroptosis-induced immune activation by disrupting lipid metabolism in TNBC cancer cells.

Prodrug-based nanomedicines  /  Ferroptosis  /  Lipid metabolism  /  Immune activation  /  Breast cancer  /  Heparin  /  Combinational therapy  /  ROS-responsiveness
Ling Lin, Zaixiang Fang, Guohao Liu, Yiwei Liu, Zhiqian Li, Dayi Pan, Yunkun Li, Hemi Kang, Xiaoding Shen, Jingyao Zhang, Qiyong Gong, Kui Luo, Jing Jing. Prodrug-based combinational nanomedicine remodels lipid metabolism for reinforced ferroptosis and immune activation[J]. Acta Pharmaceutica Sinica B, 2025 , 15 (5) : 2746 -2763 . DOI: 10.1016/j.apsb.2025.03.016
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1.　Introduction
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Ferroptosis, also known as “iron death”, is a form of programmed cell death, and it could be a promising strategy for cancer therapy1,2. Ferroptosis inducers have been reported to suppress tumor growth, reshape the tumor microenvironment, and enhance immunotherapy responses to overcome resistance to conventional therapies in various tumors3,4. Treatment with ferroptosis inducers is imperative for aggressive tumors that are difficult to treat, such as triple-negative breast cancer (TNBC)5,6. Beyond its direct cytotoxic effect, ferroptosis can promote immunogenic cell death (ICD) and invigorate the immune system by inducing the release of damage-associated molecular patterns (DAMPs), presenting antigens to dendritic cells (DCs) to form mature ones, activating immune cells proliferation7. The immune activation process, stimulated by adaptive immune response, could lead to recognition and attack of tumor cells8. It is particularly important in “cold” tumors, including TNBC, which have a low level of immunogenicity and poor immune cell infiltration9,10.
Although TNBC cells are susceptible to ferroptosis compared to conventional chemotherapies, several challenges exist to inducing ferroptosis and destroying tumors effectively11,12. Ferroptosis is triggered by overwhelmed lipid oxidation, which is influenced by lipid metabolic properties and is governed by intracellular redox balance13. Multiple intracellular reductive systems, represented by the glutathione peroxidase 4 (GSH/GPX4) pathway, could defend against oxidative stress and maintain redox balance14. Attacking one therapeutic target by affecting certain antioxidants or intervening in the metabolism might not sufficiently disrupt the redox balance, resulting in insufficient lipid oxidation and incomplete ferroptosis in tumor tissues15. The remaining tumor cells may recover and migrate into the circulation, leading to tumor migration and invasion16,17. Therefore, combination therapies with two or more therapeutic agents may simultaneously modulate metabolism, increase oxidative lipid levels, and exert synergistic pharmacological actions to enhance ferroptosis.
Paclitaxel (PTX), a first-line chemotherapeutic agent for treating TNBC, is known for disrupting microtubule dynamics and inhibiting cell mitosis18. It has also been shown to modulate metabolic processes and induce ferroptosis when combined with other therapeutic modalities, thereby increasing the anticancer sensitivity of these combination treatment modalities against TNBC cells19,20. In addition, pyropheophorbide-a (Ppa), a photodynamic therapy (PDT) agent, generates reactive oxygen species (ROS) upon light irradiation, effectively exerting oxidative stress in tumor cells, which is a critical trigger of ferroptosis21,22. Therefore, combining Ppa and PTX may be a promising synergistic approach to induce ferroptosis in TNBC cells and activate adaptive immune responses in TNBC tissues.
Nevertheless, the combinational use of small molecular compounds for clinical application is constrained by several issues, including low solubility, poor tumor targeting, and high systemic toxicity23,24. For these concerns, novel biomaterials have been developed to serve as efficient drug carriers for delivering Ppa and/or PTX25-27. Although natural polysaccharides such as heparin (HP) have advantageous biological properties, they are inherently inadequate as stand-alone drug carriers28,29. Consequently, enhancing the functionality of natural polysaccharide through modification has emerged as a reliable strategy. One strategy is to graft hydrophilic and biocompatible polymers onto HP, effectively transforming HP into a versatile nanocarrier for co-delivery of multiple therapeutic agents30,31. Poly(N-(2′-hydroxy) propyl methacrylamide) (pHPMA) is a hydrophilic polymer with high biocompatibility and has been widely investigated as a substituent group in a drug delivery carrier32. This approach not only improves the solubility and bioavailability of therapeutic agents but also enhances their targeting efficiency to tumor tissues, thus facilitating the co-delivery of drugs in a combination nanomedicine for tumor treatment33.
In this study, we grafted pHPMA onto HP to produce HP-pHPMA (HH) via reversible addition–fragmentation chain transfer (RAFT) polymerization. HH was employed to deliver a photosensitizer, Ppa, to yield HP-pHPMA-Ppa (HH-Ppa) and an anti-tumor drug, PTX, to yield HP-pHPMA-PTX (HH-PTX). The co-assembly of HH-PTX and HH-Ppa resulted in forming a prodrug-based combinational nanomedicine, HH-PP (Scheme 1). Upon penetration into tumor tissues and accumulation in tumor cells, ROS generated by Ppa in HH-PP under laser irradiation-induced cell death and promoted the release of PTX. HH-PP was also found to significantly affect lipid metabolism, as evidenced by decreased GSH and GPX4 and increased levels of oxidized phosphatidylethanolamine (oxPE) and other oxidative lipids. The disruption of lipid homeostasis markedly enhanced ferroptosis in tumor cells, thereby activating an adaptive immune response. HH-PP treatment yielded an encouraging tumor growth inhibition rate in a tumor-bearing animal model.
2.　Materials and methods
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2.1.　 Materials, cell lines and animals
All chemicals used were of analytical grade, and their sources were listed in Supporting Information 1.1. Instruments were shown in Supporting Information 1.2. Trackers and probes, except specifically mentioned ones, were obtained from Beyotime Biotechnology (Chengdu, China), and antibodies were purchased from BD Pharmingen (Shanghai, China). BODIPY 581/591 C11 were obtained from Jiqi Biotechnology (Shanghai, China). Cells were purchased from the Chinese Academy of Science Cell Bank (Shanghai, China). Mice and nude mice (female, 6–8 weeks) were purchased from Gem Pharmatech (Chengdu, China). All experimental procedures were executed according to the protocols approved by the Animal Ethics Committee of West China Hospital of Sichuan University (No. 20220602003).
2.2.　 Synthesis and characterization of compounds and polymers
The synthesis routes are shown in Supporting Information 1.3. Briefly, the macromolecular chain transfer agent, HP-CTA, was synthesized from HP-DBCO and CTA-N3. HP-CTA, together with N-(2′-hydroxyl)-propyl methyl propylene amide (HPMA), and 2-propenamide, 2-methyl-N-[2-(2-pyridinyldithio)ethyl] (MA–SS–Py) were used to form HH which was subsequently employed to produce HP-pHPMA-SH (HH-SH). HH-SH was introduced to form HH-Ppa and HH-PTX when it reacted with Mal-Ppa and Mal-CA-TA-PTX, respectively. Reaction products were characterized via the following measurements, including NMR spectra, HRMS, LC–MS analysis, GPC analysis, DSC analysis, or FTIR analysis, and details were exhibited in Supporting Information 1.4.
2.3.　 Preparation and characterization of nanoparticles
HH-PTX and HH-Ppa were dissolved in deionized water and prepared using an ultrasound machine. Co-assembled HH-PP from HH-PTX and HH-Ppa at different ratios were prepared by physically mixing HH-Ppa and HH-PTX at a high concentration of 100 mg/mL, dispersing them in deionized water and then subjecting to ultrasonic treatment. For HH-PTX imaging, 5-carboxy-x-rhodamine(Rox)-labeled HH-PTX was prepared, named RoxHH-PTX, and its co-assembled HH-PP was renamed RoxHH-PP.
Characterizations of the nanoparticles include size and zeta potential measurements, morphology observation, in vitro stability and photostability evaluation, and in vitro singlet oxygen production assessment. General measurement methods and instruments were listed in Supporting Information 1.5.
2.4.　 Cellular uptake
Mouse breast cancer cells (4T1 cells) were cultured in Roswell Park Memorial Institute (RPMI-1640) with 10% fetal bovine serum (FBS) (v/v), 100 U/mL penicillin G, and 100 μg/mL streptomycin at 37 °C under 5% CO2. Cells (2 × 103) were treated with different nanoparticles for 24 h (RoxHH-PTX, HH-Ppa, RoxHH-PP) (Ppa: 1 μg/mL, and PTX: 1.4 μg/mL). After incubating with cell organelle-specific trackers and staining with Hoechst 33,342, fluorescence images of organelles, Rox, Ppa, and cell nuclei were captured by confocal laser scanning microscopy (CLSM).
2.5.　 Cell toxicity studies
In vitro experiments were conducted in the following groups: Ppa and PTX without irradiation (Ppa + PTX), free Ppa and free PTX with irradiation (Ppa + PTX + L), HH-PTX, HH-Ppa, HH-Ppa with irradiation (HH-Ppa + L), HH-PP, and HH-PP with irradiation (HH-PP + L). 4T1 cells (2 × 104) were co-cultured with different nanoparticles for 24 h (Ppa: 2 μg/mL, and PTX: 2.8 μg/mL). Subsequently, the HH-Ppa + L and HH-PP + L groups underwent irradiation (1 J/cm2, 660 nm, 16 mW/cm2, 62 s) and then incubated for another 6 h. After incubation, cells were sampled for the following analyses. For live and dead viability, cells were treated with a live and dead cell viability kit, and their images were captured under a microscope. Cells were fixed with 4% polyformaldehyde and incubated with Hoechst 33,342 and actin and tubulin trackers to assess the impact of nanoparticles on microtubules and microfilaments. Cellular microtubule and microfilament morphologies were captured under CLSM. For assessing intracellular ROS generation and measuring the mitochondria membrane potential, cells were co-cultured with 2,7-dichlorodihydrofluorescein diacetate (DCFH-DA), and 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC-1), respectively. The fluorescence images of ROS or JC-1 were acquired with an inverted fluorescence microscope or CLSM. Cells were harvested for semi-quantification analysis with flow cytometry.
2.6.　 TEM imaging
4T1 cells (1 × 108) were treated with HH-Ppa, HH-PTX or HH-PP for 24 h (Ppa: 2 μg/mL, and PTX: 2.8 μg/mL), and the cells in the HH-Ppa + L and HH-PP + L groups were irradiated under 660 nm laser (1 J/cm2, 16 mW/cm2, 62 s), following with incubation for another 1 h. Cells were collected and fixed with 2.5% glutaraldehyde. Subsequently, they were sectioned by a slicer to obtain bio-TEM images.
2.7.　 Lipid oxidation analysis and ferroptosis-related assays
4T1 cells (2 × 105) were subjected to the same treatment in 2.5. The concentrations of malondialdehyde (MDA), GSH, and lactate dehydrogenase (LDH) were assessed using an MDA kit, a GSH assay kit, and an LDH assay kit, respectively. The results were normalized based on the cell number according to the manufacturer's instructions. Cells were stained with BODIPY 581/591 C11 for lipid oxidation analysis and Hoechst 33,342. Cells were harvested for semi-quantification analysis with flow cytometry, and their images were acquired using CLSM.
2.8.　 Untargeted lipidomic study
4T1 cells (1 × 1010) were treated similarly in 2.6. Lipids were collected and prepared according to the previous report34. Instrument parameters were shown in Supporting Information 1.8. The raw data were acquired on HPLC–MS/MS, and lipids were extracted and identified via MS-DIAL software (n = 5).
2.9.　 Mouse model construction and in vivo biodistribution
Xenograft tumors were generated by subcutaneous injection of 4T1 cells (50 μL, 1 × 107/mL) into the right posterior backs of the Balb/c nude mice. Tumor-bearing mice were randomized into five groups (Saline, Ppa + Rox, RoxHH-PTX, HH-Ppa, RoxHH-PP; Ppa: 5 mg/kg, Rox: 0.49 mg/kg; n = 4). When the tumor size reached 100 mm3, these mice were injected with different formulations via the tail vein and subjected to living imaging. At 72 h post-injection, their organs were collected for ex-vivo imaging after sacrifice. Frozen sections of tumors were prepared, and fluorescence images were captured using CLSM.
2.10.　 Pharmacokinetics analysis
Female Balb/c mice were randomly divided into four groups, and they were intravenously injected with Ppa + Rox, RoxHH-PTX, HH-Ppa, or RoxHH-PP (Ppa: 5 mg/kg, Rox: 0.49 mg/kg; n = 4). 20 μL of blood samples from the orbital venous plexus were collected at different time points post-injection. The blood samples were added with 80 μL of H2O and stored at 4 °C for 24 h. The samples were centrifuged at 3500 rpm for 15 min, and the supernatants were extracted with 300 μL of DMSO. The Ppa and Rox concentrations in the supernatant were measured via a microplate reader (Ex: 570 nm, Em: 620 nm for Rox, Ex: 640 nm, Em: 680 nm for Ppa).
2.11.　 In vitro ICD effect
4T1 cells (1 × 105) were subjected to the same treatment in 2.5. Cells were stained sequentially with high mobility group box 1 (HMGB1) and Hoechst 33,342. For calreticulin (CRT) staining, these cells were fixed and then stained with an actin tracker and a CRT probe.
2.12.　 Stimulation of BMDCs maturation
Mouse bone marrow cells collected from mice tibia and femur were cultured in the RPMI medium supplemented with 20 ng/mL human granulocyte-macrophage colony-stimulating factor (GM-CSF). They were stimulated to generate bone marrow DCs (BMDCs) by culturing for 6 days. After 4T1 cells were processed with the same treatment in 2.5, the spent medium from 4T1 cells was added into the BMDCs medium for BMDCs maturation stimulation for 24 h. The level of DC maturation was investigated with flow cytometry by assessing the portion of CD80+CD86+ cells in CD11c+ cells (n = 3).
2.13.　 In vivo anti-tumor experiments and biosafety evaluation
Tumor-bearing Balb/c mice were randomized into eight groups, and they received intravenous injections of different formulations presented in 2.5 via tail vein on Days 1 and 9 (Ppa: 5 mg/kg, PTX: 7 mg/kg; n = 5). After 24 h post-injection, tumors in the Ppa + PTX + L, HH-Ppa + L, HH-PP + L groups were irradiated (96 J/cm2, 660 nm, 320 mW/cm2, 5 min). The tumor volume and the mice's body weight were monitored twice daily. Euthanasia was performed if the tumor length exceeded 15 mm or the mice's body weight decreased by more than 20%. After 17 days of treatment, tumors were harvested, and tumor growth inhibition (TGI) rates were calculated from the equation, as shown in Eq. (1):
TGI=100–[mt/mc×100]
where mt is the tumor weight from the treatment group, and mc is the tumor weight from the control group. The mice's blood was withdrawn for biochemistry analysis. The main organs were stained with H&E, and tumor tissues were subjected to immunohistochemistry (Ki67 and CD31) and immunofluorescence analysis (TUNEL).
2.14.　 In vivo immune activation study and Oil Red O staining
Tumor tissues from 2.13 were subjected to immunofluorescence analysis (HMGB1, CRT, CD4, and CD8). They were frozenly sectioned, and the tumor frozen slices were stained with Oil Red O after fixing with 10% formaldehyde and rinsing with 60% isopropanol.
2.15.　 Statistical analysis
Statistical analyses were performed, including one-way ANOVA, unpaired t-test, or Tukey's multiple comparison test, using GraphPad Prism 10 software. Data are presented as mean ± standard deviation (SD, ns, not significant. ∗P < 0.05. ∗∗P < 0.01. ∗∗∗P < 0.001.). Principal component analysis (PCA) of lipidomes data was performed using the SIMCA 16.0 software (Sartorius AG, Göttingen, Germany). Heat mapping, volcano analysis, and fold change analysis were performed using MetaboAnalyst 4.0 (McGill University, Quebec, Canada).
3.　Results and discussion
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3.1.　 Synthesis and characterization
HP was selected as the base polymer to prepare the combinational nanomedicine due to its inherent biocompatibility and versatility for chemical modification. However, native HP has a limited capacity as a drug carrier, and we modified it with pHPMA via RAFT polymerization. This modification allows precise control of the molecular weight of the drug carrier and enables the conjugation of therapeutic agents such as Ppa and PTX onto the carrier. As shown in Supporting Information Scheme S1, HH-Ppa and HH-PTX were synthesized through condensation, polymerization, and click reaction (Supporting Information Figs. S1–S9). HP-CTA, a macromolecular chain transfer agent, was synthesized in two steps. In the first step, HP was modified with DBCO-NH2 to create HP-DBCO according to a previously reported method35. Differential scanning calorimetry (DSC) revealed a distinct exothermic peak for HP-DBCO at 229.33 °C, differing from that of native HP (245.56 °C), DBCO-NH2 (an endothermic peak, 102.83 °C) and a mixture of HP and DBCO-NH2 (244.83 °C), confirming successful conjugation (Fig. 1A). UV spectra further supported the modification with absorption peaks at 294 and 312 nm for HP-DBCO that were not present in native HP (Supporting Information Fig. S10). Quantitative analysis showed that the content of DBCO-NH2 in HP-DBCO was 12.68%, which was calculated from a standard curve (Supporting Information Fig. S11).
Subsequently, HP-CTA was synthesized using alkyne and azide groups via a copper-free click reaction. 1H nuclear magnetic resonance (1H NMR) spectra confirmed the presence of aromatic protons at 8.0–7.0 ppm, characteristic peaks of the DBCO and CTA groups (Figs. S5–S6). In contrast, Fourier-transform infrared spectra (FTIR) showed the disappearance of the azide peak of CTA-N3 at 2229.43 cm−1, verifying successful synthesis (Supporting Information Fig. S12). DSC analysis displayed an exothermic peak at 228.00 °C for HP-CTA, indicating a slight shift from HP-DBCO (Fig. 1A). UV absorbance analysis also confirmed a peak at 307 nm, consistent with the spectral properties of CTA (Fig. 1B). The content of CTA in HP-CTA was measured at 9% (Supporting Information Fig. S13).
HH was synthesized by RAFT polymerization using HP-CTA as the chain transfer agent, HPMA as the monomer, and 2,2′-azobis[2-(2-imidazolin-2-yl) propane] dihydrochloride as an initiator under an argon atmosphere. Subsequently, HH-Ppa and HH-PTX were synthesized by conjugating Mal-Ppa36 and Mal-TA-CA-PTX37, respectively, to HH through a thiol-maleimide click reaction. The characterization by gel permeation chromatography revealed a slight increase in the molecular weight (Mw) for HH-Ppa and HH-PTX in comparison to HH, which indicated successful conjugation of the drug (Supporting Information Fig. S14, Supporting Information Table S1). 1H NMR analysis confirmed the presence of characteristic peaks for Ppa and PTX, verifying successful synthesis of HH-Ppa and HH-PTX (Figs. S7–S9). The drug loading content (DLC) was measured to be 7.0% for Ppa and 10% for PTX (Supporting Information Fig. S15). To map the distribution of HH-PTX in vitro and in vivo, we synthesized rhodamine (Rox)-labeled HH-PTX, RoxHH-PTX, according to a previous method37, with a Rox content of 0.7%.
The self-assembly and co-assembly behaviors were investigated, showing the critical aggregation concentrations of HH-Ppa and HH-PTX were determined to be 3.69 μg/mL and 12.23 μg/mL, respectively, indicating that both could form nanoparticles in an aqueous solution (Supporting Information Table S2). Alterations in size, polydispersity index (PDI), and correlation coefficients of HH-Ppa and HH-PTX at varying concentrations via dynamic light scattering (DLS) indicated that both HH-Ppa and HH-PTX were capable of maintaining a nanoscale size at a low concentration (Supporting Information Fig. S16, Supporting Information Tables S3 and S4). The particle size and PDI of the co-assembled product formed from HH-Ppa and HH-PTX were further investigated at varying HH-PTX/HH-Ppa ratios. The results showed that a 1:1 mass ratio of HH-PTX to HH-Ppa resulted in an optimal particle size and PDI, and the resulting product, HH-PP (DLC of PTX: 5%; DLC of Ppa: 3.5%), was used for the following in vitro and in vivo studies (Supporting Information Fig. S17, Supporting Information Table S5). The hydrodynamic diameters of HH-Ppa and HH-PTX were 114.2 ± 6.7 and 150.6 ± 12.2 nm, respectively. In contrast, the diameter of the co-assembled HH-PP decreased to 106.4 ± 1.6 nm (Table S5). The zeta potentials of HH-Ppa, HH-PTX, and HH-PP were −14.0 ± 1.0, −24.6 ± 1.7, and −19.5 ± 1.3 mV, respectively, indicating that three polymers could circulate in the blood without engulfment by the reticuloendothelial system (Supporting Information Table S6). Notably, the resulting HH-PP exhibited a uniform spherical morphology as confirmed by transmission electron microscopy (TEM), which was different from the nanofiber morphology of HH-Ppa and an irregular shape of HH-PTX (Fig. 1C). These results indicate that HH-PP was not a physical mixture, but a co-assembled product. This stable co-assembly configuration was essential for effectively co-delivery of Ppa and PTX and their synergistic action within tumor cells.
This co-assembly technology could not only coordinate the action of the two drugs in tumor cells but also improve their stability under storage conditions and in a physiological environment. The stability results demonstrated that HH-PP exhibited high stability with negligible alterations in the particle size or PDI in H2O, PBS, and DMEM with 10% FBS over 120 h, thereby supporting prolonged circulation and preventing premature release (Fig. 1D and E, Supporting Information Fig. S18). In addition, photostability analysis confirmed that co-assembly improved the photostability of the photosensitizer in different solutions. HH-PP exhibited a 5.3% loss in the absorbance in PBS with 0.1% SDS and a 7.4% loss in DMSO, indicating a slower degradation rate compared to HH-Ppa alone (Fig. 1F). Furthermore, we evaluated the ability of HH-Ppa to generate singlet oxygen, a critical ROS for the induction of ferroptosis38, using a singlet oxygen sensor green (SOSG) reagent. The results showed that HH-Ppa and HH-PP produced a significantly elevated level of singlet oxygen under laser irradiation compared with Ppa when dissolved in PBS (Fig. 1G (1)). Additionally, the fluorescence intensity of SOSG was significantly enhanced when the assembly structures of HH-Ppa and HH-PP were disrupted in the presence of SDS. (Fig. 1G (2)). Overall, these findings demonstrated that our rationally designed HH-PP was a stable co-assembly nanomedicine for co-delivering Ppa and PTX, which could facilitate their tumor accumulation and enhance their synergistic effects on amplifying ferroptosis and the overall anti-tumor efficacy.
3.2.　 In vivo penetration and in vitro infiltration
The metabolic clearance in the circulation system was evaluated by pharmacokinetic analysis of HH-Ppa, RoxHH-PTX, RoxHH-PP, free Ppa, and free Rox. RoxHH-PP prolonged the circulation time of HH-Ppa and free Ppa. The half-life time (t1/2) of Ppa in RoxHH-PP was 15.6- and 24.9-fold longer than that in HH-Ppa and free Ppa, respectively. The area under the curve (AUC) of RoxHH-PP was 1.8- and 6.8-fold larger than that of HH-Ppa and free Ppa. RoxHH-PP slightly prolonged the circulation time of RoxHH-PTX. The t1/2 was 72.34 h in RoxHH-PP and 59.17 h in RoxHH-PTX. The AUC of RoxHH-PP was 1.4-fold larger than that of RoxHH-PTX (Fig. 2A and B, Supporting Information Tables S7 and S8). Next, the in vivo distribution of HH-Ppa, RoxHH-PTX, RoxHH-PP was observed with living imaging. HH-Ppa, RoxHH-PTX, and RoxHH-PP were injected into 4T1 tumor-bearing nude mice through tails, and Rox and Ppa signals were monitored at different pre-set time points (Supporting Information Fig. S19). Compared to RoxHH-PTX and HH-Ppa, RoxHH-PP displayed stronger fluorescence signals of Rox and Ppa in vivo at all time points (Fig. 2C and Supporting Information Fig. S20A). In contrast to the faint fluorescence signal observed in the free Ppa group at 24 h post-injection, HH-Ppa and RoxHH-PP displayed strong fluorescence signals at 72 h post-injection (Fig. S20A). Both pharmacokinetic analysis and live imaging results suggested RoxHH-PP had a prolonged circulation duration compared to HH-Ppa, RoxHH-PTX, and free Ppa.
The mice tumors and organs were collected at the end of the living imaging experiment and followed with ex vivo fluorescence imaging to investigate the distribution of nanoparticles. Stronger fluorescence signals in tumors after treatment with RoxHH-PP compared to those treated with RoxHH-PTX and HH-Ppa (Fig. 2D and E, Fig. S20B and S20C). Frozen sections of tumors were analyzed to analyze the tumor penetration ability of RoxHH-PTX, HH-Ppa, and RoxHH-PP. Results showed that both Ppa and Rox signals could be detected in both the peripheries and central zones of the tumors after treatment with RoxHH-PP, which confirmed that RoxHH-PP could penetrate deeply into the tumor tissue after it was intravenously injected into these mice (Fig. 2F).
The PTX concentration in the blood and the tumor tissue before laser irradiation and at different time points post-irradiation after a single injection of HH-PP was quantified by LC–MS/MS to assess the targeted release of PTX from HH-PP in the tumor tissue. The results showed that free PTX in the serum exhibited a very low concentration, which was below five nanograms (ng) per 200 μL serum. By contrast, the PTX concentration in the tumor tissue was much higher than that in the serum. The PTX concentration was 63.02 ± 16.49 ng per 100 mg tumor tissue, and it increased to 110.52 ± 20.22 ng per 100 mg tumor tissue after laser irradiation. The PTX concentration in the tumor tissue was gradually decreased to the level before laser treatment at 24 h post-irradiation. These results illustrate that HH-PP could be stable in the circulation system. PTX could be released in tumor tissues, and laser irradiation efficiently promoted its release from HH-PP in the tumor tissue (Supporting Information Fig. S21).
To further assess the penetration ability of RoxHH-PP in cancer cells, multicellular spheroids (MCSs) from 4T1 cells were prepared and precultured with RoxHH-PP before CLSM analysis. These results shown in Supporting Information Fig. S22 confirmed there were strong fluorescence signals of Ppa and Rox from the periphery to the inner zone of MCSs, which further supported deep penetration of HH-PP within tumor tissues. Additionally, we evaluated the cellular uptake of RoxHH-PP by 4T1 cells and its intracellular distribution in organelles. 4T1 cells were co-cultured with RoxHH-PP, and the fluorescence signals of Rox in RoxHH-PTX and RoxHH-PP, as well as Ppa in HH-Ppa and RoxHH-PP, were captured using CLSM at different time points. The results in Supporting Information Fig. S23A showed that fluorescence signals of Ppa and Rox could be detectable at hour 2, and their signal intensity was gradually enhanced as the incubation time extended. The fluorescence signals were semi-quantified via flow cytometry after internalization by 4T1 cells to compare the cellular uptake levels of different nanoformulations. Both CLSM images and flow cytometry results suggested the fluorescence signal of either Ppa or Rox was statistically intensified in 4T1 cells co-cultured with RoxHH-PP compared to those without any treatment. However, there was no statistical difference in the Ppa signal intensity in the cells after uptake of HH-Ppa and RoxHH-PP, nor the Rox signal intensity in the cells after uptake of RoxHH-PTX and RoxHH-PP (Fig. 2G and H, Fig. S23B), suggesting co-assembled formulation, RoxHH-PP, could be internalized by 4T1 cells at a similar level as RoxHH-PTX and HH-Ppa. Intracellular distribution of RoxHH-PP in organelles was further revealed. Colocalization of the signal from organelle trackers and the signals of Rox and Ppa in RoxHH-PP was captured using CLSM. The CLSM images supported that there was a wide intracellular distribution of RoxHH-PP after its internalization, and it was seen in organelles, including mitochondria, lysosomes, and the endoplasmic reticulum (ER) (Fig. 2I and J, Supporting Information Figs. S24–S26). All these results suggested that RoxHH-PP had a remarkable extension in its circulation time, a pronounced penetration ability in tumor tissues, a high level of cellular uptake, and a wide distribution in organelles after internalization, and these attributes of RoxHH-PP may fundamentally contribute to its potent anti-tumor effects.
3.3.　 In vitro anticancer potency
To assess the synergistic effects of HH-PP, the anti-tumor efficacy of HH-PP against 4T1 cells was investigated. First, the cytotoxicity of HH-PP at different ratios of HH-PTX to HH-Ppa was evaluated using CCK-8 assays to calculate the combination index (CI) via the CompuSyn software. The optimal synergism of HH-PP under irradiation was observed at a mass ratio of HH-PTX to HH-Ppa of 1:1, displaying the lowest CI in the fraction affected (Fa) range of 0.15–0.85 (Supporting Information Fig. S27). This was in accordance with the results of particle size and PDI of HH-PP (Fig. S17, Table S5). Therefore, HH-PP with a mass ratio of HH-PTX to HH-Ppa of 1:1 was used for the following studies. Cell death staining was applied for cytotoxicity of the nanoformulations against 4T1 cells. A higher percentage of dead cells indicated by positive PI staining appeared in the HH-PP and laser irradiation treated group (HH-PP + L) compared to HH-PTX and HH-Ppa + L group (Fig. 3A). A commonly used method to assess cytotoxicity via an apoptosis kit was then applied. There were no positively stained cells in the early apoptosis quadrant after incubation with HH-PP (Supporting Information Fig. S28). These treated cells were aggregated, which may be due to disturbed cell membranes after treatment with HH-PP + L, and apoptotic cells were barely detected in flow cytometry plots. The results suggest that 4T1 cell death after the treatment of HH-PP + L may not be due to apoptosis.
To unveil the predominant inducing mechanism for synergistic cytotoxicity exerted by HH-PP + L, the anti-tumor efficacy of Ppa and PTX against 4T1 cells was assessed separately. Morphological changes in cellular microtubules and microfilaments after 24-h treatment with HH-PTX or HH-PP were observed. The images revealed a similar level of shrinkage in the HH-PTX and HH-PP groups. The uniform distribution of tubulin and actin in the plasma was seen in the control group (Fig. 3B and Supporting Information Fig. S29). Cytotoxicity of Ppa has been ascribed to ROS generation under irradiation. Therefore, ROS generation was evaluated in the cells after treatment with Ppa-containing nanoformulations. Fluorescence images revealed that there were abundant cells with positive ROS signals after treatment with either HH-Ppa or HH-PP under irradiation (Fig. 3C), and flow cytometry analysis confirmed similar levels of intracellular ROS production in the cells treated with HH-Ppa and laser irradiation (HH-Ppa + L) and HH-PP + L (Fig. 3D and E). These results indicated that neither PTX nor Ppa played a dominant role in synergistic cytotoxicity in HH-PP.
Since HH-PP was distributed in different cellular organelles after internalization, the effects of HH-PP on several organelles were evaluated to demystify the mechanism(s) for synergistic cytotoxicity of HH-PP in addition to the therapeutic action of Ppa and PTX. The mitochondrial function was evaluated with a mitochondrial membrane potential kit, JC-1. JC-1 aggregates appeared in the cells without any treatment in the control group, suggesting a high mitochondrial potential in the normal mitochondria. In contrast, after culturing with different formulations for 12 h, a stronger signal of JC-1 monomers appeared in the HH-PP + L-treated group, suggesting a low mitochondria potential in the abnormal mitochondria (Fig. 3F, Supporting Information Fig. S30). An apoptosis morphology with condensation of the cytoplasm represented by a darker color of the cytoplasm and shrinkage of cells represented by a smaller diameter were observed in the TEM image of the 4T1 cells treated with HH-PTX. By contrast, 4T1 cells treated with HH-Ppa + L and HH-PP + L revealed apparently different morphologies. Furthermore, cells in the HH-PP + L-treated group with an enhancement in the ER membrane gap, suggesting these cells experienced much more severe ER stress compared to the cells in the treatment group of HH-Ppa + L (Supporting Information Fig. S31). These results implied that co-assembled HH-PP not only induced microtube and microfilament inhibition from HH-PTX and ROS generation from HH-Ppa under irradiation but also influenced the ER and mitochondrial function. All these factors could contribute to synergistic cytotoxicity of HH-PP, triggering a different cell death type instead of apoptosis.
3.4.　 Inducing of ferroptosis in tumor cells by HH-PP
HH-PP + L-treated cells revealed an abnormal morphology of mitochondria with an increase in the mitochondrial membrane density indicated by a darker color of the mitochondrial inner membrane and a reduction in the mitochondrial cristae indicated by a widened space between the mitochondrial cristae, suggesting the predominant cell death mode in these tumor cells may be ferroptosis (Fig. 4A). Severe swelling with bubbles on the plasma membranes in these cells after treatment was observed in the HH-PP + L group (Fig. 4B). Although the cytoplasmic feature of cell swelling is a typical morphology of pyroptosis, our next study on pyroptosis-associated protein expression did not support the cell death mode of pyroptosis. Specifically, we investigated the expression levels of key proteins associated with pyroptosis in 4T1 cells, including the N-terminal fragment of Gasdermin-D (GSDMD-N) and its upstream protein cleaved caspase-12,39,40. There were no statistical differences in the expression levels of GSDMD-N and cleaved caspase-1 in tumors among these treatment groups, implying pyroptosis may not be sufficiently induced in the HH-PP + L treatment group compared to other groups. No statistical difference in the expression level of cleaved caspase-3 was seen in tumor tissues between the treatment by HH-PTX and HH-PP + L, implying no enhanced apoptosis in the HH-PP + L treatment group compared to other groups (Supporting Information Fig. S32). Since the swelling appearance that was an indicator of damaged cell membranes could be observed in ferroptosis41, the abnormal morphology of the cells treated with HH-PP + L, specifically, their shape and size and their mitochondrial texture, suggested the principal cell death mode in these tumor cells may be ferroptosis.
Ferroptosis is a regulated cell death characterized by overwhelming intracellular lipid oxidation42. The most common trigger of lipid oxidation is GSH depletion accompanied by a decrease in the GPX4 expression or deactivation of GPX443. Therefore, the GPX4 expression and the GSH content were investigated to confirm ferroptosis triggered by HH-PP + L treatment. The expression of GPX4 in the tumor tissue was evaluated after mice were treated with formulations for 24 h and irradiated for another 24 h. Immunohistochemistry images of GPX4 revealed that the mice tumors treated with HH-PP + L had a lower expression of GPX4 than those in other treatment groups (Fig. 4C). The content of the intracellular antioxidant substance, GSH, was decreased after treatment with HH-PP + L compared to the control and other treatment groups (Fig. 4D). These effects induced by HH-PP + L could directly trigger lipid oxidation.
The lipid oxidation degree was then investigated. MDA, the lipid oxidation end product, was detected to confirm the level of oxidative lipids in the HH-PP + L-treated cells (Fig. 4E and Supporting Information Fig. S33). An oxidized lipid tracker, BODIPY 581/591 C11, was employed to reveal the level of lipid peroxidation (LPO) in cells. Representative CLSM images showed that the proportion of cells with LPO was remarkably increased in the HH-PP + L group compared to other treatment groups or the control group (Fig. 4F). It was also revealed that oxidation state of BODIPY 581/591 C11 staining was dispersed in the entire cytoplasm other than on the cell membrane only, suggesting a wide impact on the intracellular lipids triggered by HH-PP + L treatment (Fig. 4F). Flow cytometry analysis revealed the proportion of cells with positive LPO staining increased to 89.98% in the HH-PP + L-treated group, compared to 0.13% in the control group (Fig. 4G and Supporting Information Fig. S34). After cells experienced ferroptosis, the overwhelming intracellular lipid oxidation could eventually interrupt cell membranes (Fig. 4B). A much higher level of LDH was released from cells after exposure to HH-PP + L compared to that in the control group, suggesting the cell membrane may be damaged in the cells with HH-PP + L treatment (Fig. 4H). These results confirmed that cells treated with HH-PP + L experienced reinforced ferroptosis, evidenced by typical morphological and biochemical characteristics.
3.5.　 Ferroptosis caused by lipid disorder
Lipid oxidation is a key process in ferroptosis, and it is highly dependent on lipid compositions44. To reveal the impact of the treatment with HH-PP + L on lipid compositions in 4T1 cells, non-targeted lipidomics analysis was employed to evaluate the levels of intracellular lipid oxidation and lipid modulation. After 4T1 cells were treated with HH-PP + L, intracellular lipid was extracted, the lipid samples were run on LC–MS/MS, and lipids were identified via MS-DIAL. A total of 1892 lipids in different lipid categories were extracted and identified. Principal component analysis (PCA), a commonly used data analytics method for classification in lipidomic datasets, was applied and confirmed the differences between the cells treated with HH-PP + L and the control cells via the SIMCA 16.0 software (Fig. 5A). The changes in individual lipids were evaluated via a heat map. Ubiquitous variations throughout all categories of lipids were seen between HH-PP + L-treated cells and the control cells (Supporting Information Fig. S35). Specifically, 533 lipids were statistically upregulated, and 154 lipids were downregulated in the cells treated with HH-PP + L (Fig. 5B). Lipids abundance in each category was analyzed to display the degree of lipids variations in different categories, and oxidative lipids were statistically increased in the HH-PP + L-treated cells (Fig. 5C). To distinguish oxidative lipid variations, a radar plot was applied to present variations in individual types of oxidative lipids. The results showed that the most abundant species in the oxidative lipid type was oxPE, which is a main component of cell membranes. Other oxidized lipids species, including oxidized free fatty acids (FAs), oxidized phosphatidycholine (oxPC), oxPE, oxidized phosphatidylglycerol (oxPG), oxidized phosphatidylinositol (oxPI), and oxidized phosphatidylserine (oxPS), were also increased in the cells with HH-PP + L treatment compared to the cells without any treatment (Fig. 5D). Among these 111 oxidative lipids detected by LC–MS/MS, 40 of them were statistically increased in the HH-PP + L-treated cells compared to the control, and 33 of them were increased with a Log2 (FC) of greater than 1 (Supporting Information Fig. S36). Therefore, HH-PP treatment of tumor cells with irradiation resulted in the oxidation of a wide spectrum of lipids in various categories, suggesting that the oxidizing reaction of these intracellular lipids did not have an apparent preference for lipid categories and structures.
Apart from increased oxidative lipids, ubiquitous lipid disorder without structure diversity throughout all categories of lipids (Supporting Information Fig. S37) implied lipid synthesis, metabolism, transformation, and storage may be modulated after treatment with HH-PP + L. To unveil the mechanism for the modulation of the lipids in the tumor cells, we evaluated statistically significant lipid variations in different categories in the cells between the HH-PP + L group and the control group by estimating the relative abundance of lipids in different categories, especially triacylglycerol (TAG) and diacylglycerol (DAG). In cells, there is a dynamic equilibrium between these two related lipids and phospholipids (PLs). Specifically, FAs, as a substitute group of TAGs, DAGs, and PLs, could be shuttled among these lipids45. There are three FAs in the structure of TAGs, which could release one FA to generate DAGs, and they could be easily packaged for PLs synthesis. Excess PLs inside the cells could be oxidized by ROS to form oxPLs (including oxPC, oxPE, oxPG, oxPI, and oxPS). Either ether PLs and oxPLs could eventually trigger ferroptosis. On the contrary, the storage of FAs in TAGs could reduce the FAs pool and inhibit PLs formation, which could lead to a reduction in membrane lipids oxidation. It is noted that there was a remarkable increase in DAGs and PLs, but a decrease in TAGs, implying HH-PP + L triggered ferroptosis in cells through decreasing TAGs and increasing PLs (Fig. 5C).
TAGs and other categories of neutral lipids, such as CLs, are stored in lipid droplet (LD) to serve as a source for antioxidation46. We then accessed LD content in the cells after treatment with HH-PP + L to evaluate TAG storage. LDs could be observed in the control cells in the Oil Red O staining images and TEM images. They were barely seen in the cells after treatment with HH-PP + L, suggesting the TAG storage vesicles were reduced by the intervention of HH-PP + L (Fig. 5E and F). Furthermore, the cells treated with HH-PP + L experienced ER stress may also inhibit lipid synthesis, resulting in decreased TAG content and LD breakdown (Fig. S31). Therefore, intervention with HH-PP + L could increase lipid oxidation through LD breakdown and TAG degradation, eventually inducing ferroptosis (Fig. 5G, Supporting Information Fig. S38).
3.6.　 ICD induced by ferroptosis
During ferroptosis, the adaptive immune response system, which plays a dominant role in immune surveillance and immune attack of tumor cells to suppress tumor progression, could be activated47. To be specific, tumor cell membranes are disrupted by lipid oxidation to release DAMPs. HMGB1 released from cell nuclei and CRT exposed to cell membranes are the most significant DAMPs, which can induce DCs maturation and promote immune cell proliferation. The adaptive immunoreaction response could eventually suppress tumor proliferation48. We first studied released DAMPs, including HMGB1 and CRT, to assess adaptive immunoreaction activation. HMGB1 is primarily located in the nucleus, and it acts as a companion for DNA in a normal situation. It can also be shuttled to the cytoplasm as a typical DAMP. CRT is a major calcium-binding protein, and it is normally located in the ER. It can be overexpressed and secreted to cell membranes to enhance cell immunogenicity. As shown in Fig. 6A, the level of HMGB1 in the nucleus decreased after treatment with HH-PP + L, resulting in its translocation to the cytoplasm and subsequent release into the extracellular space. CRT was uniformly distributed in the cytoplasm, and it was also exposed to the cell membrane surface in the treatment group with HH-PP + L. The signal of CRT was the strongest in the HH-PP + L-treated group. Therefore, ferroptosis induced ICD in tumor cells to release DAMPs, which sent an “eat me” signal to the immune surveillance system for antigen presentation.
Activation of DCs maturation after the stimulation of tumor cells treated with HH-PP + L was assessed in vitro. Fig. 6B illustrates the protocol for the cultivation and activation of BMDCs. In brief, bone marrow cells were harvested, and their differentiation into BMDCs was stimulated. Meanwhile, 4T1 cells were co-cultured with HH-PTX, HH-Ppa, and HH-PP for 24 h and another 12 h after irradiation. The spent media from 4T1 cells were added into the cell culture media of BMDCs for 24 h cultivation. The level of DCs maturation was analyzed via flow cytometry by evaluating the portion of matured DCs, which were cells with CD80+CD86+ in CD11c+ DCs. The portion of mature DCs was 17.85% in BMDCs in the control group. In comparison, it was increased to 38.65% in BMDCs co-cultured with the spent media after 4T1 cells were treated with HH-PP + L, and this was statistically higher than that in BMDCs that were co-incubated with the spent media after treatment of tumor cells with HH-PTX and HH-Ppa + L (Fig. 6C and D). Therefore, HH-PP + L treatment of tumor cells could activate DCs maturation and induce adaptive anti-tumor immunity responses.
DAMPs release and immune cell proliferation were also studied in vivo. Mice were intravenously injected with different formulations through tail veins, and the tumor tissues in PTX + Ppa + L, HH-Ppa + L, and HH-PP + L treatment groups were exposed to laser for irradiation at 24 h post-injection. DAMPs expression levels and immune cell densities in tumor tissues were detected via immunofluorescence staining at 48 h post-injection. The levels of HMGB1 and CRT were elevated in the HH-PP + L-treated group compared to those in other groups (Fig. 6E–G). T cells, the major type of immune cells related to adaptive immune responses, were investigated. Among these T cells, CD8+ cytotoxic T cells (CTLs) have been considered as the anti-tumor immune effector cells for attacking tumor cells. CTLs can produce perforin, granzymes, and cytokines to induce tumor cell death but keep nonmalignant cells intact. CD4+ T helper (Th) cells are reported to provide cytokine-mediated support for CTLs expansion and their function. It was observed that CTLs proliferated, and their infiltration level was increased in the tumor tissues in the HH-PP + L treatment group compared to the other groups. There were no significant differences in the Th cells number among all experimental groups (Fig. 6E). Thus, the ratio of CD8+ T cells to CD4+ T cells was remarkably boosted in the HH-PP + L-treated group compared to other treatment groups (Fig. 6H). Therefore, the anti-tumor efficacy of HH-PP was strengthened through enhancing immunogenic responses, supported by DAMPs release, DCs maturation, and CTLs proliferation, eventually suppressing tumor progression.
3.7.　 In vivo anticancer effects of HH-PP
To evaluate the in vivo anti-tumor effect of HH-PP, tumor-bearing mice were grouped and intravenously injected with different nanoformulations (Fig. 7A). There were two injections on Days 1 and 9, and irradiation was implemented on Days 2 and 10 in the groups treated with Ppa-containing nanoformulations. During the entire course, the mice tumor volumes were monitored every other day, and they were persistently increased in the control group. In contrast, the tumor growth was restrained after the first injection of HH-PP and irradiation. The therapeutic effect of HH-PP was maintained until the experiment was completed (Fig. 7B). Although the mice tumor volumes were reduced to some degrees in other treatment groups compared to the mice injected with saline as a control, the degree of reduction in tumor volumes in these treatment groups was lower than that in the HH-PP + L-treated group. The tumor volume was used to determine the endpoint of the animal experiment. Mice were anesthetized and executed when the tumor volume in the mice injected with saline grew to larger than 1500 mm3. At the end of the experiment, tumors were harvested for measuring the tumor weight and calculating the TGI. The tumor weight was 0.21 ± 0.02 g in the HH-PP + L-treated group, while it was 1.57 ± 0.11 g in the control group (Fig. 7C, Supporting Information Table S9). The TGI in the treatment groups varied from 38.89% to 86.63%, and the highest TGI was seen after treatment with HH-PP + L (Fig. 7D). These results confirmed synergistic anti-tumor effects of HH-PP + L (Fig. 7C–E).
Anti-tumor potency was further evaluated in vivo through tumor immunohistochemistry studies, including cell death, cell proliferation, and tumor angiogenesis. TUNEL staining was widely applied to various cell death modes, such as pyroptosis and ferroptosis, although it was initially invented for apoptosis investigation. The images confirmed a large number of TUNEL-positive cells or dead cells in the HH-PP + L-treated group. The signal of Ki67, a cell proliferation marker, in the tumor tissue suggested that cell proliferation was suppressed in the groups treated with HH-Ppa + L, HH-PP, and HH-PP + L. The highest level of restraining tumor cell proliferation was seen in the HH-PP + L-treated group. The images of CD31, a marker of new blood vessel formation, confirmed the lowest density of blood vessels in the tumor tissue after treatment with HH-PP + L (Fig. 7F and Supporting Information Fig. S39). All these results confirmed that HH-PP + L could exert an excellent anti-tumor effect in vivo.
The safety of these nanoformulations was also evaluated both in vitro and in vivo. The hemolytic assay was conducted by mixing 10% of red blood cells in PBS and different concentrations of nanoformulations. The hemolysis rates of HH-PTX and HH-Ppa were lower than 4%, and that of the assembled nanoformulations HH-PP were lower than 1% (Supporting Information Fig. S40 and Supporting Information Table S10). The mice's body weight was monitored during the entire experimental period to analyze the systemic toxicity of HH-PP, and the results showed there were no statistical differences in the body weight among all experimental groups (Supporting Information Fig. S41A). Besides, the mice's serum was collected for blood chemistry tests. The levels of the biochemistry parameters, including alanine transaminase (ALT), aspartate transaminase (AST), creatine kinase (CK), creatinine (CREA), and UREA in the peripheral blood of the treated mice were equivalent to those in the control mice, indicating that the treatment with HH-PP had a negligible effect on the function of the liver, heart, and kidney (Fig. 7G). The main organs of the mice were harvested, weighed, and stained with H&E at the end of the experiment. There were no statistical differences in the weights of main organs except spleen in all treatment groups compared to the control group (Fig. S41B–S41E). The spleens of tumor-bearing mice were enlarged, but they returned to a normal size following treatment with HH-PP + L (Fig. S41F), suggesting that HH-PP + L treatment may improve the overall health of the mice after tumor suppression. H&E staining images revealed no visible abnormal morphologies in all treatment groups (Supporting Information Fig. S42). Therefore, HH-PP has a great biosafety profile and can be readily transitioned into clinical studies.
4.　Conclusions
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We successfully synthesized two heparin-based graft polymers (HH-PTX and HH-Ppa) through RAFT polymerization and co-assembled them into HH-PP as a combinational nanomedicine for delivering Ppa and PTX. HH-PP displayed excellent stability in the simulated physiological environment in vitro. The pharmacokinetic analysis showed that RoxHH-PP prolonged the circulation time and increased the AUC of HH-Ppa and RoxHH-PTX. The live in-vivo images supported that HH-PP exhibited pronounced targeting of tumor cells and enhanced infiltration in tumor tissues. Following treatment with HH-PP and laser irradiation, ferroptosis was induced in tumor cells by disrupting lipid homeostasis, as evidenced by significant oxidation of lipid metabolites. Notably, the lipid peroxidation was amplified by the breakdown of LDs and TAG degradation, which further enhanced ferroptosis. After ferroptosis of tumor cells triggered by HH-PP + L, its anti-tumor effect was reinforced by activating immune responses, including DAMPs release, DCs maturation, and CTL proliferation. Importantly, HH-PP displayed a high biosafety profile by performing hemolytic assays and biochemistry tests, monitoring the mice's body weight, and staining major organs after harvest. Therefore, a novel and promising strategy has been demonstrated to induce ferroptosis of tumor cells via targeting the vulnerability of lipid metabolism by an intelligent drug delivery system, and its anti-tumor effect could be enhanced through immunology activation.
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Year 2025 volume 15 Issue 5
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doi: 10.1016/j.apsb.2025.03.016
  • Receive Date：2024-12-09
  • Online Date：2026-09-17
Article Data
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  • Received：2024-12-09
  • Revised：2025-02-15
  • Accepted：2025-02-20
Affiliations
    aDepartment of Radiology, Huaxi MR Research Center (HMRRC), Institution of Radiology and Medical Imaging, Breast Center, Institute of Breast Health Medicine, State Key Laboratory of Biotherapy, Core Facilities, West China Hospital, Sichuan University, Chengdu 610041, China
    bCollege of Chemical Engineering, Sichuan University, Chengdu 610041, China
    cFunctional and Molecular Imaging Key Laboratory of Sichuan Province, NHC Key Laboratory of Transplant Engineering and Immunology, Research Unit of Psychoradiology, Chinese Academy of Medical Sciences, Chengdu 610041, China
    dXiamen Key Lab of Psychoradiology and Neuromodulation, Department of Radiology, West China Xiamen Hospital of Sichuan University, Xiamen 361021, China

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表12种不同金属材料的力学参数
科
Family		 属数
Number of
genus		 种数
Number of
species		 占总种数比例
Percentage of
total species (%)		 属
Genus		 种数
Number of
species		 占总种数比例
Percentage of total
species (%)
鹅膏菌科Amanitaceae 2 11 5.26 鹅膏菌属 Amanita 10 4.78
小菇科 Mycenaceae 2 12 5.74 丝盖伞属 Inocybe 5 2.39
多孔菌科 Polyporaceae 8 14 6.70 蜡蘑属 Laccaria 5 2.39
红菇科 Russulaceae 3 23 11.00 小皮伞属 Marasmius 6 2.87
小菇属 Mycena 11 5.26
光柄菇属 Pluteus 5 2.39
红菇属 Russula 17 8.13
栓菌属 Trametes 5 2.39
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