Double coordination-driven Fe–DOX synergy enhances ferroptosis and immunogenic cancer therapy

Abstract

Ferroptosis, an iron-dependent form of cell death driven by lipid peroxidation, has emerged as a potential therapeutic strategy for cancer. However, intrinsic resistance, off-target toxicity, and poor redox efficiency limit the effectiveness of existing ferroptosis-inducing platforms. This study reports novel heparin-derived metal coordination nanoparticles (HDMC NP) that leverage catechol–metal–drug coordination chemistry to enhance ferroptosis and immunogenic cell death (ICD). In this self-assembled supramolecular system, doxorubicin (DOX) directly coordinates to Fe³⁺ centers established by catechol–Fe³⁺ interactions on the heparin backbone, forming a stable HDMC NP with high drug-loading efficiency. This coordination not only stabilizes the complex under neutral conditions but also triggers disassembly in response to pH changes in the tumor microenvironment. Notably, the DOX–Fe³⁺ pairing supports sustained redox activity, amplifying ROS production and ferroptotic stress in a coordination-dependent manner beyond the effect of either component alone. In animal models, HDMC NP suppresses tumor growth by inducing both ferroptosis and ICD, characterized by calreticulin exposure and CD8⁺ T cell infiltration. Collectively, this platform provides a redox-active nanotherapeutic based on coordination-defined assembly with potential for broad applications in metal-ion modulated diseases.

Graphical abstract

A catechol–metal–drug supramolecular platform enables pH-triggered ferroptosis and immune-activated tumor cell death, presenting a modular, carrier-free strategy for redox-based cancer immunotherapy.

1. Introduction

Cancer is one of the leading causes of mortality worldwide. Despite advances in chemotherapy, radiotherapy, and immunotherapy, therapeutic resistance and off-target toxicity continue to undermine treatment efficacy1, 2, 3, 4, 5. The tumor microenvironment (TME), characterized by hypoxia, oxidative stress, and dysregulated iron metabolism, has been increasingly recognized as a key contributor to therapeutic resistance5, 6, 7, 8. Ferroptosis, a distinct iron-dependent non-apoptotic cell death pathway driven by lipid peroxidation9, 10, 11, has recently emerged as a promising strategy for overcoming resistance12, 13, 14, 15, 16. Unlike conventional apoptosis, iron-catalyzed lipid peroxidation initiates ferroptosis, resulting in oxidative damage and preferential vulnerability of cancer cells^13,17. However, many existing ferroptosis-inducing systems exhibit poor structural stability, burst drug release, and systemic toxicity^18,19. Fe-based therapies often require highly acidic conditions to generate sufficient reactive oxygen (ROS), resulting in suboptimal efficacy within the mildly acidic TME^20,21.

Polysaccharides and iron alone are insufficient to induce immunogenic ferroptosis; thus, ferroptosis-inducing agents are required. Doxorubicin (DOX), a widely used anthracycline chemotherapeutic agent, exhibits antitumor activity primarily by intercalating DNA and inhibiting topoisomerase II, leading to DNA damage and apoptosis22, 23, 24. Beyond its canonical apoptotic mechanism, DOX interacts with iron ions (Fe³⁺/Fe²⁺)25, 26, 27 to enhance ROS production, thereby triggering ferroptosis28, 29, 30. This synergistic effect arises from a redox interaction in which DOX reduces Fe³⁺ to Fe²⁺. This reduction enables Fe²⁺ to react with endogenous hydrogen peroxide through the Fenton reaction, producing highly cytotoxic hydroxyl radicals (·OH)^31,32. Recent studies further indicate that nuclear DNA damage and the DNA damage response (DDR), including ATM/ATR signaling, modulate iron metabolism and redox homeostasis, thereby sensitizing cancer cells to iron-dependent oxidative stress33, 34, 35, 36. Such redox cycling amplifies lipid peroxidation and ferroptotic stress, establishing the DOX–Fe pairing as an efficient inducer of ferroptosis^27,37. Notably, although similar DOX–Fe redox chemistry is implicated in cardiotoxicity, it may represent a potent therapeutic mechanism when selectively harnessed in cancer cells38, 39, 40, 41. Tumor cells frequently exhibit dysregulated iron metabolism, elevated basal ROS levels, and compromised redox homeostasis, rendering them markedly more susceptible to sustained oxidative stress42, 43, 44, 45. Under these conditions, DOX–Fe-driven ROS amplification and lipid peroxidation can be selectively exploited to induce ferroptotic cell death, thereby transforming a liability in normal tissues into a therapeutic advantage in cancer^46,47. This highlights the necessity of a delivery system that spatially and temporally restricts DOX–Fe redox activity to the tumor microenvironment while preserving its ferroptosis-inducing potential.

To overcome these limitations, we introduce a novel therapeutic design based on catechol–metal–drug coordination chemistry. This approach leverages the strong affinity between catechol groups and metal ions, such as Fe³⁺, Cu²⁺, and Zn²⁺, to directly assemble small-molecule drugs into stable supramolecular complexes48, 49, 50. This coordination-based strategy conceptually resembles metal–organic frameworks (MOFs) because both involve metal–ligand interactions to construct ordered nanostructures51, 52, 53, 54. However, unlike conventional MOFs that require rigid crystalline scaffolds and exogenous linkers55, 56, 57, the system developed in this study forms soft, biocompatible, and supramolecular nanoparticle (NP) in which the drug itself functions as a building block. This minimal coordination network provides several intrinsic advantages: (i) disassembly in an acidic TME due to pH-responsiveness; (ii) delivery of chemically unmodified, intact drug molecules; and (iii) versatility, as it can be extended to other therapeutic agents and metal ions with compatible coordination sites. This biologically adaptive supramolecular design offers a promising platform for delivering redox-responsive drugs to cancer cells and other inflammation-associated diseases.

In this study, we developed a pH-responsive heparin-derived metal coordination nanoparticle (HDMC NP) as a model for implementing this strategy (Scheme 1). In the HDMC system, DOX formed a stable coordination with Fe³⁺ through catechol moieties on the heparin backbone, resulting in a supramolecular nanostructure with high drug-loading efficiency. Heparin contributes to aqueous solubility and biocompatibility, while enhancing coordination stability through its negatively charged sulfate and carboxylate groups58, 59, 60, 61. The resulting HDMC NP exhibited greater stability under physiologically relevant near-neutral conditions; however, they disassembled in an acidic TME, enabling localized drug release. Notably, the coordination between DOX and Fe³⁺ served both as a structural element and a catalytic engine for ferroptosis. The DOX–Fe³⁺ complex facilitated redox cycling and amplified ROS production, resulting in enhanced lipid peroxidation and ferroptotic stress. This synergistic effect significantly enhanced cytotoxicity compared with DOX or Fe³⁺ alone. This system promoted immunogenic cell death (ICD), as evidenced by increased calreticulin (CRT) exposure and dendritic cell activation. Through this integrated design, HDMC NP enables coordinated drug delivery, ferroptosis activation, and immune stimulation within the TME.

Scheme 1.

Schematic representation of the self-assembled metal–polysaccharide coordination system for ferroptosis-driven synergistic cancer therapy. Heparin–dopamine conjugate (UDOPA) coordinates with Fe³⁺ ions and doxorubicin (DOX) via metal–catechol interactions to form self-assembled supramolecular nanoparticles (HDMC NP). The system harnesses DOX–Fe³⁺ redox synergy to amplify ferroptosis via Fenton-type ROS generation and lipid peroxidation. Simultaneously, oxidative stress promotes ICD and dendritic cell activation, stimulating antitumor immunity. Supramolecular design ensures tumor-specific activation and addresses the key limitations of conventional ferroptosis inducers, such as burst release, instability, and systemic toxicity.

2. Materials and methods

2.1. Materials

Antibiotic antimycotic solution (100X), acetonitrile (ACN), anhydrous dimethyl sulfoxide (DMSO), deuterium oxide (D₂O), dimethyl sulfoxide-d₆, DispaseⅡ, DNaseⅠ, dopamine hydrochloride, Dulbecco's modified Eagle's medium (DMEM), 2,7-dichlorodihydrofluorescein diacetate (DCF-DA), 1-ethyl-3-(3-dimethyl aminopropyl)carbodiimide (EDC), ethanol, heparin sodium salt from porcine intestinal mucosa (UFH), iron (Ⅲ) chloride, N-hydroxysuccinimide (NHS), 2-(N-morpholino) ethane sulfonic acid monohydrate (MES monohydrate), methanol, phosphate-buffered saline (PBS), tetramethylsilane (TMS), trifluoroacetic acid (TFA), and 2-(4-amidinophenyl)-6-indolecarbamidine dihydrochloride (DAPI) were purchased from Sigma–Aldrich (St. Louis, MO, USA). BODIPY 581/591 C11 (Lipid Peroxidation Sensor), CellTracker Deep Red-C34565, CellTracker Green CMFDA-C7025, Collagenase TypeⅡ powder, and Foxp3 transcription factor-staining buffer kit were purchased from Thermo Fisher Scientific (Waltham, MA, USA). FerroOrange was purchased from DOJINDO Laboratories (Kumamoto, Japan). Spectra/Por 7 Dialysis Membrane Pre-treated RC Tubing was purchased from Repligen (Waltham, MA, USA). Doxorubicin hydrochloride was purchased from FuthreCHEM (Seoul, Republic of Korea). Roswell Park Memorial Institute (RPMI) 1640 was purchased from WELGENE (Gyeongsan-si, Republic of Korea). Fetal bovine serum (FBS) was acquired from Gibco (Thermo Fisher Scientific, Waltham, MA, USA). Cell culture SPLInsert-35224 and Cell Strainer-93040 were purchased from SPL Life Sciences (Pocheon-si, Gyeonggi-do, Republic of Korea). The EZ-cytox kit was obtained from DoGenBio (Seoul, Republic of Korea). Neutral buffered formalin (10%) was purchased from HuBenTech (Damyang, Republic of Korea).

2.2. Experimental animals

Five-week-old male and female C57BL/6 mice were purchased from Orient Bio (Seongnam, Republic of Korea). The animals were housed in an animal facility under a 12 h light–dark cycle, and the temperature and humidity were maintained at 25 ± 1 °C and 50 ± 5%, respectively. To ensure animal welfare, CO₂ euthanasia was performed using a gradual-fill method (30%–50% chamber volume per minute) to minimize distress, in accordance with the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals. The procedure was carried out under controlled conditions with continuous monitoring, and all efforts were made to minimize pain and discomfort during the process. All animal experiments were conducted in accordance with the standard regulations of the Institutional Animal Care and Use Committee (IACUC) of Konkuk University (Ref. No. KU22078-1).

2.3. Synthesis and characterization of UDOPA

The Synthesis between various ratios of unfractionated heparin (UFH) and dopamine hydrochloride (DOPA) (1:10, 1:20, 1:30, and 1:40, molar ratio) was carried out in a 10 mmol/L MES buffer. First, activation of UFH is required. According to a UFH-DOPA (UDOPA) ratio of 1:10, UFH (100 mg) is dissolved in 10 mL of MES buffer along with EDC (38 mg) and NHS (23 mg), and the solution was stirred at 96×g in a water bath for 10 min. Then, after adding DOPA (25.3 mg), the solution was stirred at 96×g in an ice bath for 6 h. After 6 h, the synthesized material was lyophilized for 1 day to obtain a white powdered form. Additionally, UDOPA was precipitated in 25 mL of ethanol and purified by centrifugation at 1224×g, 4 °C for 4 min, repeated 4 times. The purified product was confirmed to be purified through thin-layer chromatography (TLC).

To verify the successful synthesis of UDOPA, the final product was analyzed by ¹H NMR spectroscopy using a 500 MHz spectrometer (JNM-ECZ500R/S1, JEOL, Tokyo, Japan). Samples were prepared at a concentration of 10 mg/mL in a solvent mixture of D₂O and DMSO-d₆ (2:8, v/v).

To quantify the amount of conjugated DOPA, a UV–visible spectrophotometric assay was performed using a microplate reader (SPECTROstar Nano, BMG Labtech, Ortenberg, Germany). A standard calibration curve was generated using free DOPA solutions, with absorbance measured at 280 nm in a quartz cuvette. UDOPA samples were dissolved in distilled water at 1 mg/mL, and their absorbance at 280 nm was recorded. The DOPA content in each conjugate was determined based on the calibration curve, enabling comparison of conjugation efficiencies across different feed ratios (UFH: DOPA = 1:10, 1:20, 1:30, and 1:40, molar ratio). An average molecular weight of 15,000 Da was used for UFH in all calculations.

2.4. Synthesis and characterization of UDFe

UDFe were synthesized by mixing UDOPA and ferric chloride (FeCl₃) at various mass ratios (UDOPA:FeCl₃ = 20:1, 20:2, 20:4, and 20:6, w/w) in distilled water (DW), which resulted in an acidic reaction environment due to FeCl₃ hydrolysis. At an UDFe ratio of 1:5, UDOPA (100 mg) was dissolved in DW (50 mL), and the mixture was stirred at 96×g in an ice bath for 6 h to facilitate coordination. After reaction completion, the solutions were lyophilized for 24 h to obtain light green powders. The crude products were further purified by dialysis against distilled water for 48 h using regenerated cellulose membrane tubing (Spectra/Por 7, 3.5 kDa, Thermo Scientific, Waltham, MA, USA).

To determine the optimal formulation, each UDFe complex was evaluated for colloidal stability at a concentration of 0.5 mg/mL in distilled water over various time points (0, 3, 6, 12, 24, and 48 h). Following centrifugation, the supernatant was collected, and the absorbance at 334 nm—corresponding to Fe³⁺—was measured using a microplate reader (SPECTROstar Nano, BMG Labtech, Ortenberg, Germany). In addition, visual inspection was performed using plastic cuvettes containing UDFe (1 mg/mL), UDOPA (1 mg/mL), and FeCl₃ (0.1 mg/mL) in DW. To further assess coordination characteristics beyond colloidal stability, the optimal ratio was additionally evaluated by UV–Vis spectroscopy, X-ray photoelectron spectroscopy (XPS), and inductively coupled plasma optical emission spectrometry (ICP–OES, Avio 550, PerkinElmer, Waltham, MA, USA), providing complementary information on coordination-related optical features and iron binding capacity, respectively.

2.5. In silico modeling and molecular dynamics simulation

The molecular structures of iron ion (Fe³⁺), dopamine (DOPA) and doxorubicin (DOX) were generated and visualized using ChemDraw Professional 20.1.1.125 (PerkinElmer, Waltham, MA, USA). The heparin structure was obtained from the PDB database (PDB ID: 3IRJ), while UDOPA was generated by modifying heparin using BIOVIA Discovery Studio 2022 (BIOVIA, San Diego, CA, USA) to reflect its conjugation with dopamine.

For Molecular Dynamics (MD) simulations, all molecules were converted using the CHARMM force field and optimized using the Prepare Ligand protocol in Discovery Studio 2022. For pH-responsive modeling, two sets of protonation states were prepared representing mildly acidic (pH 5.5) and neutral (pH 7.2) environments. The Standard Dynamics Cascade protocol was employed to conduct the simulation, with all processes performed under NVE ensembles using the distance-dependent dielectric solvent model. To investigate whether a Fe-mediated DOPA–DOX coordination complex could form, a ternary system consisting of DOPA, Fe³⁺, and DOX was simulated for 100 ps under each pH condition. The formation of stable coordination complexes was analyzed at the final frame.

A representative DOX–Fe–DOPA structure, extracted from the MD trajectory under neutral pH conditions, was subjected to density functional theory (DFT) calculations using ORCA 5.0.2. Geometry optimization and single-point energy calculations were performed using the B3LYP functional with D3BJ dispersion correction and the def2-SVP basis set, with TightSCF convergence criteria applied. Frontier molecular orbital (FMO) analysis was conducted to evaluate the HOMO-LUMO distribution, with orbital energies and spatial distributions calculated using Visual Molecular Dynamics (VMD), version 1.9.4 (University of Illinois Urbana-Champaign, Urbana, IL, USA). The HOMO-LUMO gap and electron density localization were used to assess the coordination-induced charge transfer characteristics. To evaluate self-assembly and coordination stability in multicomponent systems, extended MD simulations were performed on UDOPA, DOX, and Fe³⁺ mixtures up to 1000 ps under both pH conditions. Energy profiles, interaction types, and coordination persistence were analyzed to assess pH-dependent structural behavior of the assembled HDMC NP system.

2.6. Synthesis and characterization of HDMC NP

HDMC NPs were prepared by reacting UDFe with DOX in DW at various mass ratios (UDFe:DOX = 20:1, 20:3, 20:6, 20:9, and 20:12 w/w). UDFe (100 mg) and DOX (5, 15, 30, 45, and 60 mg) were dissolved in DW (50 mL), and the pH of the reaction mixture was adjusted to neutral using NaOH before stirring. The solution was then stirred at 96×g at room temperature for 6 h. The resulting solution was lyophilized overnight to yield dark purple powder. Unconjugated DOX was removed by four rounds of precipitation in methanol: ethanol (1:1, v/v) followed by centrifugation at 780×g for 4 min at 4 °C. The successful purification was confirmed by thin-layer chromatography. The optimal formulation was determined by assessing the colloidal stability of each HDMC NP sample (0.5 mg/mL in DW). The supernatant was collected at predetermined time points (0, 3, 6, 12, 24, and 48 h) and its absorbance was measured at 500 nm using a microplate reader (SPECTROstar Nano, BMG Labtech, Ortenberg, Germany). Additionally, a photographic comparison was performed using plastic cuvettes containing HDMC NP (1 mg/mL), UDFe (1 mg/mL), and DOX (1.5 mg/mL) in DW.

To evaluate pH-dependent DOX release from HDMC NP, a dialysis-based release study was performed. HDMC NP was dissolved in DW at a concentration of 40 mg/40 mL and adjusted to acidic (pH 5–6) or neutral pH using HCl or NaOH. The solution was transferred into dialysis bags with a molecular weight cutoff of 3.5 kDa (Spectra/Por7, 3.5 kDa, Thermo Scientific, Waltham, MA, USA) and immersed in the corresponding release medium under gentle stirring. At predetermined time points, aliquots of the external medium were collected and replaced with an equal volume of fresh medium. The amount of released DOX was quantified using reverse-phase high-performance liquid chromatography (RP-HPLC).

2.7. Characterization of UDOPA, UDFe, and HDMC NP

The physicochemical properties of the UDOPA, UDFe, and HDMC NP were comprehensively evaluated by analyzing the particle size and surface charge (DLS and zeta potential), morphology (TEM and SEM), chemical bonding and coordination (FT-IR, Raman, XPS, UV–Vis), and elemental composition (EDS and XRF). The hydrodynamic diameters and surface charges of the samples were measured by DLS and zeta potential analysis (Zetasizer Nano, Malvern Instruments, Malvern, UK), respectively. Before measurement, samples were prepared in DW (1 mg/mL) and ultrasonicated for 5 min at room temperature using a JAC Ultrasonic 3010 device (KODO, Seoul, Republic of Korea). The morphological characteristics of the NPs were examined using TEM (EVO MA 10, Carl Zeiss, Oberkochen, Germany) and field-emission SEM (FE-SEM; MERLIN, Carl Zeiss, Oberkochen, Germany). TEM was used to visualize the internal structure and confirm nanoparticle formation, while FE-SEM was used to analyze the surface characteristics.

The samples were analyzed by UV–Vis spectroscopy using a microplate reader (SPECTROstar Nano, BMG Labtech, Ortenberg, Germany). The electronic transitions and coordination-driven spectral shifts were determined using XPSPEAK41 software (Raymund Kwok, The Chinese University of Hong Kong, Hong Kong, China). Samples were prepared in DW (0.5 mg/mL) and analyzed to confirm the successful synthesis of the complexes, including the incorporation of DOPA into UFH and Fe-coordination-induced absorbance changes, validating nanostructural stability. Powder samples were analyzed by XPS using a PHI GENESIS spectrometer (ULVAC-PHI, Chigasaki, Japan) to further elucidate the metal coordination state, particularly the oxidation state of Fe and its interactions with the catechol moiety. The elemental composition was further analyzed by EDS (JSM-7610F, JEOL, Tokyo, Japan) and XRF (ZSX Primus IV, Rigaku, Tokyo, Japan) to confirm the presence of Fe in UDFe and HDMC NP, thereby validating the metal–organic coordination structure. The molecular interactions were evaluated, and the formation of coordination bonds was confirmed by analyzing powdered samples by FT-IR (Cary 610/660, Agilent Technologies, Santa Clara, CA, USA) and Raman spectroscopy (Alpha 300R, Oxford Instruments, Abingdon, UK). FT-IR spectra were recorded using KBr-pelletized samples. Raman spectroscopy provided additional structural insights into the metal–ligand interactions. This comprehensive characterization confirmed the successful formation of the metal–organic coordination structure, stability, and physicochemical properties, establishing UDOPA, UDFe, and HDMC NP as a well-defined platform for drug delivery applications.

2.8. Cell culture and in vitro cytotoxicity assay

B16F10.OVA cells, a genetically modified subline of B16F10 murine melanoma cells (RRID: CVCL_0159), were kindly provided by Prof. In-San Kim (Korea Institute of Science and Technology, KIST). These cells were engineered to stably express ovalbumin (OVA) as a model tumor-associated antigen, allowing evaluation of antigen-specific immune responses. Human primary dermal fibroblasts (HDFa; ATCC PCS-201-012; RRID not available for this primary cell type) were obtained from PromoCell (Heidelberg, Germany) and cultured according to the supplier's instructions. Cells were maintained in high-glucose Dulbecco's modified Eagle medium (DMEM, Gibco, Thermo Fisher Scientific, Waltham, MA, USA), supplemented with 10% fetal bovine serum (FBS, Gibco, Thermo Fisher Scientific, Waltham, MA, USA) and 1% antibiotic–antimycotic solution (Gibco, Thermo Fisher Scientific, Waltham, MA, USA), under standard conditions (37 °C, 5% CO₂).

For cytotoxicity analysis, the cells were seeded in a 96-well plate at a concentration of 5.0 × 10³ cells/well and preincubated at 37 °C in a 5% CO₂ incubator for 24 h. Subsequently, B16F10.OVA cells were further incubated with UFH, UDOPA, UDFe, and HDMC NP at concentrations ranging from 0.0005 to 500 μg/mL for 48 h. Free doxorubicin (DOX) cytotoxicity was evaluated separately at concentrations ranging from 0.0001 to 100 μmol/L under identical conditions. Cell viability was assessed using the EZ-Cytox cell viability assay kit (DoGenBio, Seoul, Republic of Korea) and quantified using a microplate reader (SPECTROstar Nano, BMG Labtech, Ortenberg, Germany) at 450 and 600 nm. The cell viability was calculated based on the absorbance values of the measured samples compared to those of the control group. To quantitatively evaluate the synergistic interaction between UDFe and DOX in the HDMC NP formulation, Combination Index (CI) analysis was performed using CompuSyn software (CompuSyn Inc., Paramus, NJ, USA) based on the Chou–Talalay method⁶². To evaluate cytotoxicity toward normal cells, human fibroblast cells were seeded under identical conditions and treated with UFH, UDOPA, UDFe, or HDMC NP for 24 h, followed by viability assessment using the same assay protocol. The cells were pretreated for 1 h with ferrostatin-1 (Fer-1, 2 μmol/L), deferoxamine (DFO, 100 μmol/L), or Z-VAD-FMK (20 μmol/L) prior to treatment with HDMC NP (10 μg/mL, DOX-equivalent concentration) for 48 h for inhibitor-based mechanistic studies.

2.9. Western blot assay

To evaluate the expression of GPX4 and SLC7A11 in B16F10.OVA cells, the cells were cultured in 100 mm cell culture dishes for 24 h in the presence of DOX, DOX/Fe, UDOPA, UDFe, and HDMC NP at a concentration of 400 nmol/L DOX-equivalent. For ferroptosis inhibition studies, cells were pretreated with ferrostatin-1 (Fer-1, 2 μmol/L) for 1 h prior to drug treatment and subsequently incubated with the indicated formulations under the same conditions. Following incubation, the cells were washed with PBS and lysed using CETi buffer (Translab, Daejeon, Republic of Korea). The lysates were then centrifuged at 18,000×g for 30 min at 4 °C to remove cellular debris. Protein concentration was determined using a BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). The quantified protein samples were mixed with SDS loading buffer and heated at 95 °C for 10 min. A total of 20 μg of protein was separated by 10%–15% SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and subsequently transferred onto polyvinylidene difluoride (PVDF) membranes. Their membranes were blocked in a 5% skim milk solution at room temperature for 1 h and subsequently washed three times with TBS-T buffer for 10 min per wash. The membranes were then incubated overnight at 4 °C with primary antibodies. After incubation, the membranes were washed three times with TBS-T buffer for 10 min per wash, followed by incubation with an anti-mouse IgG-HRP secondary antibody solution at room temperature for 1 h. Finally, the membranes were washed five times with TBS-T buffer for 5 min per wash, and protein bands were detected using SuperSignal West Pico Chemiluminescent Substrate (Thermo Fisher Scientific, Waltham, MA, USA). The antibodies used in this study are listed in Supporting Information Table S1.

2.10. Immunocytochemistry (ICC) detection

The expression levels of immune factors related to ferroptosis (GPX4, SLC7A11) and damage-associated molecular pattern (DAMPs) factors (CRT, HMGB1) in B16F10.OVA cells were evaluated using immunocytochemistry. Additionally, the levels of DCF-DA were assessed to confirm ROS release, and FerroOrange (Dojindo Laboratories, Kumamoto, Japan) was used to evaluate intracellular Fe²⁺ accumulation. The cells were cultured in 35 mm black confocal dishes (SPL Life Science, Pocheon-si, Republic of Korea) at a density of 1.0 × 10⁴ cells/dish for 3 h in DMEM containing 10% FBS and 1% AA. The culture media were then replaced, and the cells were treated with DOX, UDFe, and HDMC NP at a concentration of 0.1 μmol/L and incubated for 24 h. After incubation, the cells were washed with DPBS and fixed using paraformaldehyde (PFA). To reduce non-specific binding, blocking was performed with 5% BSA dissolved in DPBS for 10 min. After blocking, the immune factors were incubated overnight with primary antibodies diluted 1:500 in 5% BSA. The following day, the cells were incubated with secondary antibodies diluted 1:500 in 5% BSA for 1 h at room temperature. The nuclei of cells were stained with DAPI, and images were captured using the ECLIPSE Ti2 Series microscope (Nikon, Tokyo, Japan).

2.11. Analysis of intracellular iron accumulation

Intracellular iron accumulation was assessed using FerroOrange, a fluorescent probe specific for Fe²⁺. Cells were seeded in 12-well plates at a density of 1 × 10⁵ cells/well and treated with DOX, DOX/Fe, UDOPA, UDFe, and HDMC NP (200 nmol/L DOX-equivalent) for 24 h. For iron chelation studies, cells were pretreated with deferoxamine (DFO, 100 μmol/L) for 1 h before drug treatment and subsequently incubated under the same conditions. Following treatment, the culture medium was removed, and the cells were harvested. The cells were then stained with FerroOrange (1 μmol/L) and incubated at 37 °C for 30 min. After staining, the cells were washed twice with PBS and analyzed using flow cytometry (NovoCyte, Agilent Technologies, Santa Clara, CA, USA).

2.12. Quantification of intracellular reactive oxygen species (ROS) levels

ROS levels were measured using H2DCF-DA, a fluorescent probe for ROS detection. Cells were seeded in 12-well plates at a density of 1 × 10⁵ cells/well and incubated with DOX, UDOPA, UDFe, and HDMC NP (200 nmol/L DOX-equivalent) for 24 h. After treatment, the culture medium was removed, and the cells were harvested. The cells were then stained with H2DCF-DA (10 μmol/L) and incubated at 37 °C for 30 min. Following staining, the cells were washed twice with PBS and analyzed using flow cytometry (NovoCyte, Agilent Technologies, Santa Clara, CA, USA).

2.13. Evaluation of calreticulin expression on the cell surface

Calreticulin expression on the cell surface was analyzed using FITC-conjugated anti-mouse calreticulin antibody. Cells were seeded in 12-well plates at a density of 1 × 10⁵ cells/well and treated with DOX, DOX/Fe, UDOPA, UDFe, and HDMC NP (200 nmol/L DOX-equivalent) for 24 h. After treatment, the culture medium was removed, and the cells were harvested. The cells were then stained with FITC anti-mouse calreticulin antibody (1:100 dilution) and incubated at 4 °C for 30 min. Following staining, the cells were washed twice with FACS buffer and analyzed using flow cytometry (NovoCyte, Agilent Technologies, Santa Clara, CA, USA).

2.14. Fluorescence-based lipid peroxidation assay

Lipid peroxidation was evaluated using BODIPY 581/591, a lipid peroxidation-sensitive fluorescent probe. Cells were seeded in 12-well plates at a density of 1 × 10⁵ cells/well and treated with DOX, UDOPA, UDFe, and HDMC NP (200 nmol/L DOX-equivalent) for 24 h. After treatment, the culture medium was removed, and the cells were harvested. The cells were then stained with BODIPY 581/591 (1:1000 dilution) and incubated at 37 °C for 30 min. Following staining, the cells were washed twice with PBS and analyzed using flow cytometry (NovoCyte, Agilent Technologies, Santa Clara, CA, USA).

2.15. Phagocytosis assay

RAW264.7 macrophages were seeded in 100 mm cell culture dishes and stimulated with lipopolysaccharide (LPS, 500 ng/ml) for 24 h to induce differentiation into M1-polarized macrophages. The differentiated M1 macrophages were then stained with CellTracker Deep Red (C34565, Thermo Fisher Scientific, Waltham, MA, USA). B16F10.OVA cells were seeded in 12-well plates at a density of 1 × 10⁵ cells/well and treated with DOX, UDOPA, UDFe, and HDMC NP (200 nmol/L DOX-equivalent) for 24 h. Following treatment, the culture medium was removed, and the cells were washed with PBS before staining with CellTracker Green CMFDA (C7025, Thermo Fisher Scientific, Waltham, MA, USA). After staining, M1 macrophages and B16F10.OVA cells were co-cultured for 30 min to allow phagocytic interactions. Phagocytosis was subsequently analyzed using flow cytometry (NovoCyte, Agilent Technologies, Santa Clara, CA, USA) and fluorescence microscopy (Carl Zeiss Microscopy GmbH, Jena, Germany). In parallel, lipid peroxidation was further quantified using a colorimetric malondialdehyde (MDA) assay kit (Lipid Peroxidation (MDA) Assay Kit, Cat. No. ab118970, Abcam, Cambridge, UK) according to the manufacturer's instructions.

2.16. Migration assay

B16F10.OVA cells were seeded in 24-well plates at a density of 5 × 10⁴ cells/well and treated with DOX, UDOPA, UDFe, and HDMC NP (400 nmol/L DOX-equivalent) for 6 h. Following treatment, the culture medium was removed and replaced with fresh medium. A cell culture insert was then placed in the well, and RAW264.7 cells (1 × 10⁵ cells/insert) were added. After 6 h of co-culture, the insert was stained using the H&E Staining Kit (AB245880, Abcam, Cambridge, UK). The stained membrane was carefully removed, mounted onto a glass slide, covered with a cover slip, and examined under a microscope. For experiments involving flow cytometry, RAW264.7 cells were pre-stained with CellTracker Green CMFDA (C7025, Thermo Fisher Scientific, Waltham, MA, USA) before being added to the insert. After 24 h of incubation, RAW264.7 cells that had migrated into the culture medium were collected and analyzed using flow cytometry.

2.17. Ex vivo immune activity analysis

B16F10.OVA cells were seeded in 100 mm culture dishes at a density of 1 × 10⁶ cells/dish and treated with DOX, UDOPA, UDFe, and HDMC NP (400 nmol/L DOX-equivalent) for 24 h. Spleens were isolated from C57BL/6 mice, and red blood cells were removed using RBC lysis buffer. The remaining splenocytes were counted and prepared for co-culture. Following drug treatment, the culture medium was removed, and 1 × 10⁷ splenocytes were added to the wells for co-culture with B16F10.OVA cells for 24 h. To analyze immune cell populations, the cells were stained with a fluorescence-conjugated antibody solution at 4 °C for 20 min, washed twice with FACS buffer, and analyzed using flow cytometry.

Cell analysis was conducted as follows: T cells (CD45⁺/CD3⁺), helper T cells (CD45⁺/CD3⁺/CD4⁺), activated helper T cells (CD45⁺/CD3⁺/CD4⁺/CD25⁺), cytotoxic T cells (CD45⁺/CD3⁺/CD8⁺), activated cytotoxic T cells (CD45⁺/CD3⁺/CD8⁺/CD25⁺), NK cells (CD45⁺/NK1.1⁺), NKT cells (CD45⁺/NK1.1⁺/CD3⁺), dendritic cells (CD45⁺/CD11c⁺), activated dendritic cells (CD45⁺/CD11c⁺/CD86⁺), macrophage cells (CD45⁺/CD11b⁺/F4/80⁺), SIINFEKL expressed dendritic cells (CD45⁺/CD11c⁺/H-2Kb–SIINFEKL⁺), SIINFEKL expressed macrophage cells (CD45⁺/CD11b⁺/F4/80⁺/H-2Kb–SIINFEKL⁺)

2.18. In vivo antitumor efficacy

5-week-old male C57BL/6 mice were obtained from Orient Bio (Seongnam, Republic of Korea) and acclimatized for one week before the experiment. After the acclimatization period, B16F10.OVA cells (5.0 × 10⁵ cells in 100 μL per mouse) were subcutaneously injected into the right dorsal region. One week after tumor implantation, when the tumors were established, mice were intravenously administered saline, DOX, UDOPA, UDFe, or HDMC NP every three days for a total of 12 days. HDMC NP and DOX were administered at a dose of 2.7 mg/kg, determined based on the DOX-equivalent amount measured by absorbance at 450 nm. The body weight of drug-treated mice was measured daily using digital calipers (ASIMETO, Pleasanton, CA, USA). After 12 days of drug treatment, the mice were sacrificed, and tumors were harvested and fixed in 10% neutral buffered formaldehyde (HuBENTec, Damyang, Republic of Korea). Tumor tissues were stained with Hematoxylin and Eosin (H&E) and subjected to immunohistochemical (IHC) staining for CD8 and CRT, while apoptosis was assessed using TUNEL staining. The stained tissue sections were examined using a multimedia video microscope (OS-33DPM; OSUN HITECH, Seoul, Republic of Korea) and a digital microscope from the ECLIPSE Ti2 Series (Nikon, Tokyo, Japan). Quantification and further analysis of stained tumor tissues were performed using the ImageJ software (National Institutes of Health, Bethesda, MD, USA).

2.19. In vivo toxicity assessment

5-week-old male C57BL/6 mice were purchased from Orient Bio (Seongnam, Republic of Korea). The mice were randomly divided into three groups and intravenously administered saline, DOX, or HDMC NP at a dose of 10 mg/kg. Body weights of drug-treated mice were recorded daily for 4 days. On Day 4, blood samples were collected from the abdominal aorta for biochemical analysis. After blood collection, the mice were euthanized, and the major organs were harvested and weighed. In addition, to assess biosafety under therapeutically relevant conditions, a separate toxicity study was conducted using the same DOX-equivalent dose (2.7 mg/kg). Mice body weight changes and serum biochemical parameters were analyzed to evaluate treatment-associated systemic toxicity.

2.20. Pharmacokinetics (PK) study

Pharmacokinetic profiles were evaluated in 5-week-old male Sprague–Dawley rats following intravenous administration of Free DOX or HDMC NP at a DOX-equivalent dose of 1.35 mg/kg. At predetermined time points, whole blood was collected, and serum was obtained by centrifugation at 984×g for 15 min. Serum samples were mixed with 20% (v/v) DMSO before analysis. Free DOX levels were quantified based on their intrinsic fluorescence, whereas HDMC NP levels were determined using rhodamine B isothiocyanate (RITC) labeling using a microplate reader (SpectraMax M2, Molecular Devices, San Jose, CA, USA).

2.21. Biodistribution study

Biodistribution of HDMC NP and free DOX was evaluated in 5-week-old nude mice. B16F10.OVA cells (5 × 10⁴ cells/mouse) were subcutaneously inoculated, and tumors were allowed to grow for 1 week. The mice were then intravenously administered free DOX or HDMC NP at a DOX-equivalent dose of 2.7 mg/kg. At 48 h post-injection, the mice were euthanized, and major organs (heart, liver, spleen, lung, and kidney) and tumors were harvested. Ex vivo fluorescence signals were acquired and analyzed based on the intrinsic fluorescence of DOX using an imaging system (Visque, Vieworks, Anyang, Republic of Korea).

2.22. Single-cell RNA sequencing and bioinformatic analysis

Single-cell RNA sequencing was performed to analyze transcriptomic changes at the individual cell level. Total RNA was isolated using TRIzol reagent (Thermo Fisher Scientific, Waltham, MA, USA), and RNA quality was assessed using the TapeStation 4000 System (Agilent Technologies, Santa Clara, CA, USA). Quantification of RNA was performed with the ND-2000 Spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Libraries were prepared using the CORALL RNA-Seq V2 Library Prep Kit (LEXOGEN, Vienna, Austria) following the manufacturer's protocol. Poly(A) RNA was enriched using the Poly(A) RNA Selection Kit (LEXOGEN, Vienna, Austria), and the isolated mRNA was used for cDNA synthesis and fragmentation. Indexing was performed with Illumina indexes 1–12, and PCR amplification was used for library enrichment. The final libraries were validated using the TapeStation HS D1000 Screen Tape (Agilent Technologies, Santa Clara, CA, USA) to assess mean fragment size, and quantified using a library quantification kit on a StepOne Real-Time PCR System (Applied Biosystems, Thermo Fisher Scientific, Waltham, MA, USA). High-throughput sequencing was conducted as paired-end 100 bp reads on the NovaSeq 6000 platform (Illumina, San Diego, CA, USA). Raw sequencing data underwent quality control using FastQC, and adapter sequences and low-quality reads were trimmed with Fastp. The resulting clean reads were aligned to the reference genome using the STAR aligner, and transcript quantification was performed with Salmon. Normalization of read counts was carried out using the TMM + CPM method in EdgeR. All downstream data mining and visualization were conducted using ExDEGA software (Ebiogen Inc., Seoul, Korea).

2.23. Analysis of immune cells in tumor, TDLNs, blood, and spleen using flow cytometry following in vivo treatment

For immune profiling, mice were treated and grouped identically to those in the in vivo tumor efficacy study, receiving randomized intravenous administrations of saline, DOX, UDOPA, UDFe, or HDMC NP every 3 days for 12 days. After treatment, blood, spleen, tumor-draining lymph nodes (TDLNs), and tumors were collected and processed for flow cytometry analysis. Blood samples containing anticoagulants were subjected to red blood cell lysis using red blood cell lysis buffer, and the remaining cells were washed with fluorescence-activated cell sorting (FACS) buffer. The samples were then incubated with a fluorescently conjugated antibody solution at 4 °C for 20 min, washed twice with FACS buffer, resuspended, and analyzed using flow cytometry (NovoCyte 3000, Agilent Technologies, Santa Clara, CA, USA). The spleens were mechanically dissociated using a cell strainer (40 μm) and syringe plunger (1 mL), followed by red blood cell lysis. The samples were centrifuged at 316×g for 3 min, the supernatant was discarded, and the cells were washed with FACS buffer. The cells were stained with a fluorescent antibody solution at 4 °C for 20 min, washed twice, and resuspended for flow cytometric analysis.

2.24. TDLNs were processed using the same method as that used for the spleens

Briefly, they were mechanically dissociated using a cell strainer (40 μm) and syringe plunger (1 mL), followed by centrifugation at 316×g for 3 min. The supernatant was discarded, and the cells were washed with FACS buffer, incubated with a fluorescent antibody solution at 4 °C for 20 min, washed twice, and resuspended for flow cytometry analysis. Tumor tissue was processed by mincing tumors using scissors and enzymatically digesting in RPMI 1640 at 37 °C for 45 min using a gentleMACS Tissue Dissociator (Miltenyi Biotec, Bergisch Gladbach, Germany) with collagenase (0.2% w/v, Thermo Fisher Scientific, Waltham, MA, USA), DNase (30 U/mL, Sigma–Aldrich, St. Louis, MO, USA), and dispase (10 U/mL, Sigma–Aldrich, St. Louis, MO, USA). The resulting suspension was filtered through a cell strainer, and immune cells were isolated by density-gradient centrifugation using Histopaque-1077 (Sigma–Aldrich, St. Louis, MO, USA). The obtained cells were washed, incubated with a fluorescent antibody solution at 4 °C for 20 min, washed twice with FACS buffer, and resuspended for flow cytometric analysis. For regulatory T cell analysis, the isolated cells were fixed and stained using the Foxp3 Transcription Factor Staining Buffer Kit (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA), according to the manufacturer's instructions. Stained cells were analyzed using flow cytometry (NovoCyte 3000, Agilent Technologies, Santa Clara, CA, USA) and processed using Novoexpress software (Agilent Technologies, Santa Clara, CA, USA).

Cell analysis was conducted as follows: T cells (CD45⁺/CD3⁺), helper T cells (CD45⁺/CD3⁺/CD4⁺), activated helper T cells (CD45⁺/CD3⁺/CD4⁺/IFN-γ⁺), Foxp3⁺ T cells (CD45⁺/CD3⁺/CD4⁺/Foxp3⁺), cytotoxic T cells (CD45⁺/CD3⁺/CD8⁺), activated cytotoxic T cells (CD45⁺/CD3⁺/CD8⁺/IFN-γ⁺), PD-1⁺ T cells (CD45⁺/CD3⁺/CD8⁺/PD-1⁺), NK cells (CD45⁺/NK1.1⁺), IFN-γ-secreting NK cells (CD45⁺/NK1.1⁺/IFN-γ⁺), NKT cells (CD45⁺/NK1.1⁺/CD3⁺), IFN-γ-secreting NKT cells (CD45⁺/NK1.1⁺/CD3⁺/IFN-γ⁺), dendritic cells (CD45⁺/CD11c⁺), activated dendritic cells (CD45⁺/CD11c⁺/CD86⁺), macrophage cells (CD45⁺/CD11b⁺/F4/80⁺), M1-like tumor-associated macrophages (CD45⁺/CD11b^+/F4/80⁺/iNOS⁺), M2-like tumor-associated macrophages (CD45⁺/CD11b⁺/F4/80⁺/arginase⁺), and MDSC (CD45⁺/CD11b⁺/Gr-1⁺).

2.25. Schematic illustration

Schematic illustrations and graphical representations in this study were prepared using Adobe Illustrator, version 30.2.1 (Adobe Inc., San Jose, CA, USA).

2.26. Statistical analysis

All results are presented as mean ± standard deviation (SD) or standard error of mean (SEM), as indicated. Error bars represent the variability among independent biological replicates. Statistical comparisons between groups were performed using one-way analysis of variance (ANOVA) followed by Tukey's multiple comparisons test. All statistical analyses were carried out using GraphPad Prism 10.0 (GraphPad Software Inc., San Diego, CA, USA). Statistical significance was defined as ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001. Sample sizes were determined based on standard practice and practical considerations.

3. Results and discussion

3.1. Synthesis and characterization of UDOPA

To construct the self-assembled metal–organic coordination nanoplatform, heparin-dopamine conjugate (UDOPA) was synthesized using heparin and dopamine. Heparin, a naturally occurring polysaccharide, is a U.S. Food and Drug Administration (FDA)-approved anticoagulant with well-established biomedical applications. Dopamine, a natural catecholamine, was conjugated to heparin; the catechol moiety of dopamine can form stable coordination bonds with metal ions. Unfractionated heparin (UFH) and dopamine hydrochloride (DOPA) were conjugated at various molar ratios (UFH: DOPA = 1:10, 1:20, 1:30, and 1:40; molar ratios based on the average molecular weight of UFH) via the 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC)/N-hydroxy succinimide (NHS) coupling reaction to produce UDOPA and optimize functionalization (Fig. 1A). Based on the average molecular weight of UFH, dopamine hydrochloride was conjugated at molar ratios of 1:10, 1:20, 1:30, and 1:40. Ultraviolet–visible (UV–Vis) spectroscopy revealed the characteristic absorbance of DOPA at 280 nm, which was used to quantify its conjugation to UFH. The absorbance increased with higher DOPA feed ratios and reached saturation at a UFH: DOPA molar ratio of 1:20 (Fig. 1B). Proton nuclear magnetic resonance (¹H NMR) spectroscopy further confirmed successful dopamine conjugation, showing a characteristic DOPA peak at ∼7 ppm that was absent in native UFH, while no significant spectral changes were detected beyond the 1:20 UFH: DOPA ratio (Fig. 1C)^63,64. Fourier transform infrared (FT-IR) spectra revealed that the C O stretching peak of UFH at 1600 cm⁻¹ shifted to 1620 cm⁻¹ in UDOPA, indicating amide bond formation. The peak at 1520 cm⁻¹ corresponds to the bending vibration of the secondary amide (–NH), and the peak at 1374 cm⁻¹ is attributed to the stretching of –OH groups in the catechol moiety of dopamine, confirming successful conjugation (Fig. 1D)^63,65. The analysis of UDOPA by dynamic light scattering (DLS) revealed a particle size of 355.5 ± 55.84 nm with a polydispersity index (PDI) of 0.4226 ± 0.7141, indicating poor self-assembly and a suboptimal size distribution for further applications (Fig. 1E). The measured zeta potential of UDOPA was −45.17 ± 3.27 mV, likely due to the strong negative charge of heparin (Supporting Information Fig. S1). The transmission electron microscopy (TEM) image revealed nonuniform particle sizes, further confirming the heterogeneous morphology of UDOPA (Fig. 1F).

Figure 1.

Characterization of UDOPA and UDFe. (A) Schematic of UDOPA synthesis via the (EDC)/(NHS) coupling reaction between heparin and dopamine. (B) UV–Vis absorption spectra of UDOPA with different UFH: DOPA molar ratios of 1:10, 1:20, 1:30, and 1:40 (n = 3) at 280 nm. (C) ¹H NMR spectra of UFH, DOPA, and UDOPA confirming successful conjugation at various UFH:DOPA ratios. (D) FT-IR spectra of UFH and UDOPA. (E) Analysis of the size of UDOPA by DLS (n = 3). (F) TEM image of UDOPA (scale bar: 200 nm). (G) Schematic of UDFe synthesis via coordination of Fe³⁺ with UDOPA. At near-neutral pH, dimeric catechol–Fe coordination may occur, accompanied by a visible color change (right). (H) Macroscopic color change observed upon mixing UDOPA with FeCl₃. (I) Stability assessment of UDFe at different UDOPA:FeCl₃ ratios (n = 3). Representative photographs show pH-dependent color changes of the formulations under near-neutral conditions. (J) UV–Vis absorption spectra (∼760 nm) of UDFe at different UDOPA:FeCl₃ ratios. (K) Analysis of the size of UDFe by DLS (n = 3). (L) TEM image of UDFe (scale bar: 100 nm). XPS analysis: (M) full survey spectrum, (N) Fe 2p₃ region, and (O) O 1s spectrum, confirming coordination bonding between the DOPA moieties of UDOPA and Fe³⁺ ions. (P) UV–Vis spectra of UFH, DOPA, FeCl₃, UDOPA, and UDFe. Data are presented as mean ± SD. Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, and ∗∗∗P < 0.001.

3.2. Synthesis and characterization of UDFe

The UDFe bioconjugate was synthesized by mixing UDOPA and FeCl₃ in different mass ratios (UDOPA:FeCl₃ = 20:1, 20:2, 20:4, and 20:6, w/w) (Fig. 1G). The coordination reaction was carried out in acidic distilled water, under which Fe³⁺ preferentially forms stable 1:1 coordination with catechol groups of DOPA. The formation of the UDFe was accompanied by a visible color change, as colorless UDOPA and pale yellow FeCl₃ produced a green solution upon mixing, indicative of predominant 1:1 catechol–Fe³⁺ coordination (Fig. 1H).

The optimal UDOPA:FeCl₃ ratio was determined by evaluating the colloidal stability of UDFe particles at a concentration of 0.5 mg/mL in distilled water (DW) (Fig. 1I). UDOPA and FeCl₃ were mixed at different mass ratios and incubated at room temperature. The supernatant was collected at specific time points, and the UV–Vis absorbance was measured at 334 nm to monitor the sedimentation behavior of the UDFe particles. Among the tested formulations, UDFe prepared at a UDOPA:FeCl₃ ratio of 20:2 exhibited the highest colloidal stability, with minimal aggregation and precipitation. Notably, under near-neutral conditions and at higher catechol availability, additional catechol ligands can further coordinate with Fe³⁺, leading to ligand exchange and higher-order coordination. Consistent with this observation, UV–Vis analysis revealed a characteristic absorption band at ∼760 nm⁶⁶, which increased with Fe addition and reached saturation at the 20:2 ratio, indicating the predominance of 1:1 DOPA–Fe coordination (Fig. 1J and Supporting Information Fig. S2). This was further supported by Fe 2p X-ray photoelectron spectroscopy (XPS) analysis, which showed the largest relative satellite peak area for the 20:2 formulation (Supporting Information Fig. S3). Quantitative iron-binding analysis using inductively coupled plasma optical emission spectrometry (ICP–OES) showed that Fe incorporation reached a plateau at the UDOPA:FeCl₃ ratios of 20:2, and 20:4 (corresponding to ∼100 mg Fe per kg of UDOPA), indicating saturation of the DOPA–Fe coordination capacity (Supporting Information Fig. S4). Taken together, UDOPA:FeCl₃ ratio of 20:2 corresponds to a coordination-saturated state in which Fe³⁺ is predominantly bound to catechol groups without the presence of excess or non-coordinated iron, thereby providing the optimal balance between defined coordination stoichiometry and colloidal stability.

The size of the UDFe particle was 264.07 ± 17.06 nm with a PDI of 0.4397 ± 0.0857 (Fig. 1K). Although the PDI value of UDFe at the 20:2 ratio indicates moderate dispersibility, it should be noted that UDFe represents an intermediate coordination precursor rather than the final nanoparticle formulation. At this stage, the selection of the 20:2 ratio was primarily guided by achieving defined coordination stoichiometry and colloidal stability without excess free iron, rather than by optimizing particle dispersibility. The measured zeta potential of UDFe was −30.63 ± 0.07 mV, reflecting a reduction in the magnitude of surface negative charge compared with UDOPA, attributable to the incorporation of Fe³⁺ ions (Supporting Information Fig. S5). The TEM image of UDFe particles revealed a spherical structure at the individual particle level (Fig. 1L). In addition, quantitative particle size distribution analysis based on multiple TEM images was performed and presented as a TEM-derived size histogram, confirming that the observed morphology is representative of the overall UDFe particle population (Supporting Information Fig. S6). The apparent discrepancy between the hydrodynamic size measured by DLS (Fig. 1K) and the particle size observed by TEM (Fig. 1L) can be attributed to the supramolecular and polymer-based nature of UDFe. During TEM sample preparation, including drying and vacuum conditions, soft supramolecular assemblies may undergo partial structural rearrangement or collapse. Such differences between solution-state and dry-state measurements are commonly observed for soft supramolecular assemblies67, 68, 69.

The coordination interactions between DOPA and Fe in UDFe were further investigated by X-ray photoelectron spectroscopy (XPS), which confirmed the presence of the Fe 2p₃ peak in the UDFe spectra (Fig. 1M and Supporting Information Fig. S7). The Fe 2p₃ XPS spectrum revealed both Fe³⁺ 2p₃/₂ and Fe³⁺ 2p₁/₂ peaks, which are characteristic of FeCl₃^70,71. Importantly, deconvolution of the Fe 2p₃/₂ region revealed a distinct Fe³⁺ satellite peak, indicative of electronic transitions of Fe³⁺ ions, thereby further supporting successful coordination bonding (Fig. 1N). Moreover, the O 1s XPS spectrum of UDFe exhibited an additional Fe–O signal distinct from the characteristic heparin-associated oxygen peaks, further confirming the coordination interaction between Fe³⁺ and UDOPA (Fig. 1O)⁷². UV–Vis spectroscopy confirmed the successful synthesis of UDFe. When comparing UDOPA, FeCl₃, and UDFe, UDFe showed a significantly higher intensity and a broader peak than FeCl₃ alone, suggesting the successful coordination of Fe³⁺ into the UDOPA structure (Fig. 1P).

3.3. In silico modeling and theoretical validation of the DOPA–Fe–DOX coordination structure

Molecular dynamics (MD) simulations were conducted to examine the feasibility of DOPA–Fe–DOX coordination under different pH conditions. At neutral pH, DOPA and DOX formed a stable coordination complex with Fe³⁺, maintaining a metal–ligand framework with Fe³⁺ as the central ion (DOPA–Fe–DOX), whereas under mildly acidic conditions the coordination was disrupted, indicating that complex formation is pH-dependent, consistent with coordination behavior observed for other divalent metal ions and metal-chelating drugs (Supporting Information Fig. S8 and S9). A representative DOPA–Fe–DOX structure was observed in the 100-ps snapshot (Fig. 2A).

Figure 2.

Computational analysis of the DOPA–Fe–DOX coordination complex and self-assembly of HDMC NP. (A) MD simulation under neutral and acidic pH. (B) DFT-based optimization of the Fe–O coordination structure. (C) Frontier orbital analysis (HOMO and LUMO). (D) MD simulation (1000 ps) of the HDMC NP. (E, F) Energy profiles under different pH conditions. (G) Analysis of interaction types. (H) Quantification of molecular states of DOPA, DOX, and Fe³⁺ in MD simulations.

Density functional theory (DFT) calculations were performed on the MD-extracted structure using ORCA (B3LYP-D3BJ/def2-SVP, TightSCF) to evaluate electronic and structural stability. The self-consistent field (SCF) procedure converged after 73 cycles, yielding a single-point energy of −3706.98 Eh (Fig. 2B). The Fe–O coordination bond length ranged from 2.0 to 2.3 Å, consistent with stable coordination bonds^73,74. Mulliken charge analysis indicated partial charge delocalization around the Fe center, suggesting coordination-induced electronic redistribution. Frontier molecular orbitals were visualized using Visual Molecular Dynamics (VMD)⁷⁵. The highest occupied molecular orbital (HOMO) was mainly localized on the Fe center and coordinating oxygen atoms, whereas the lowest unoccupied molecular orbital (LUMO) was distributed over the aromatic system of DOX. The HOMO-LUMO gap of 2.23 eV reflects moderate electronic stability, further supporting the structural feasibility of the coordination complex (Fig. 2C). This coordination strategy is generalizable to other drugs, as demonstrated with curcumin (Supporting Information Fig. S10).

3.4. Molecular dynamics analysis of pH-dependent HDMC NP self-assembly

The multicomponent self-assembly behavior was examined in greater detail through an extended MD simulation lasting 1000 ps on a mixture containing UDOPA, Fe³⁺, and DOX (Fig. 2D), which constitutes the core coordination-driven building unit of the pH-responsive heparin-derived metal coordination nanoparticle (HDMC NP) system. The resulting nanocomplexes exhibited robust self-assembly through sustained Fe–ligand interactions, forming a stable network with minimal disruption that maintained its structure beyond 100 ps. These interactions remained stable throughout the simulation, maintaining the DOPA–Fe–DOX coordination structure. In contrast, under mildly acidic conditions, several DOX molecules dissociated from Fe³⁺, and only partial interactions with DOPA persisted, indicating instability of the coordination complex in low-pH environments (Supporting Information Fig. S11).

The total and potential energies were significantly lower under neutral conditions than under acidic conditions, indicating a thermodynamically more favorable assembly (Fig. 2E and F). Quantitative interaction analysis of the MD trajectories revealed a higher number of metal–acceptor and electrostatic interactions at neutral pH, whereas hydrogen bonding was favored under acidic conditions, leading to disruption of coordination (Fig. 2G). In addition, direct counting of molecular states showed that the proportion of unbound DOX or DOPA molecules was higher under acidic conditions, while formation of the DOPA–Fe–DOX complex occurred exclusively at neutral pH. These results indicate that mildly acidic conditions destabilize metal–ligand coordination, whereas neutral conditions support robust supramolecular assembly through Fe³⁺ coordination with DOPA and DOX, thereby driving the formation of stable HDMC nanoparticles (Fig. 2H).

3.5. Preparation and characterization of HDMC NP

pH-responsive heparin-derived metal coordination nanoparticle (HDMC NP) was synthesized under near-neutral conditions by coordinating UDFe with DOX to form the characteristic DOPA–Fe–DOX structure (Fig. 3A). Under neutral conditions, Fe³⁺ can coordinate with two ligands, and the quinone-hydroquinone moiety in DOX enables metal coordination, facilitating complex formation76, 77, 78. The color changes from green (UDFe) and red (DOX) to dark purple upon the formation of HDMC NP further confirms the coordination interactions (Fig. 3B). Although DOX contains additional potential coordination sites, such as hydroxyl and amino groups on the daunosamine moiety, their contribution to HDMC NP assembly appears to be minor under the present conditions. Under near-neutral conditions, the DOX amino group predominantly exists in a protonated (NH₃⁺) form^79,80, as confirmed by N 1s XPS analysis, and MD simulations showed only rare involvement (∼4.13%) in metal coordination. Accordingly, HDMC NP assembly is mainly governed by DOPA–Fe coordination and the redox-active quinone moiety of DOX (Supporting Information Fig. S12).

Figure 3.

Structural characterization and pH-responsive behavior of HDMC NP. (A) Schematic of the formation of the DOPA–Fe–DOX complex via the coordination of UDFe with DOX. (B) Photographs showing the color change from green (UDFe) and red (DOX) to dark purple upon HDMC NP formation. (C) Stability analysis at different UDFe:DOX mass ratios (n = 3). (D) Size analysis by DLS and SEM image (n = 3). (E) Zeta potential measurement of HDMC NP (n = 3). (F) TEM image (scale bar: 200 nm). (G) Raman spectra of the UDOPA/DOX physical mixture and HDMC NP. XPS analysis: (H) full survey spectrum, (I) Fe 2p₃ region, and (J) O 1s spectrum. (K) UV–Vis spectra of HDMC NP, UDFe, and DOX. (L) pH-responsive behavior analyzed by UV–Vis spectroscopy. (M) DOX release monitored by UV–Vis spectroscopy and visual color change at different pH levels (n = 3). (N) DLS analysis of HDMC NP: stable nanoparticles at neutral pH (left) and disrupted self-assembly with DOX release at mildly acidic pH (right). Data are presented as mean ± SD. Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, and ∗∗∗P < 0.001.

The formulation of HDMC NP was optimized by combining UDFe and DOX at mass ratios of 20:1, 20:3, 20:6, 20:9, and 20:12; the 20:3 ratio was identified as the optimal ratio based on the stability test (Fig. 3C). The analysis of size distribution of HDMC NP by DLS revealed a uniform size of 214.57 ± 2.92 nm with a PDI of 0.2456 ± 0.0103, indicating a well-distributed NP population, consistent with scanning electron microscopy (SEM) results (Fig. 3D). Compared with UDFe, which serves as a coordination precursor, the formation of HDMC NP resulted in markedly improved particle dispersibility and colloidal stability. The measured zeta potential of HDMC NP was −26.69 ± 0.43 mV, indicating a stable formulation (Fig. 3E). The TEM image confirmed that the HDMC NP displayed spherical morphology and uniform dispersion (Fig. 3F). In addition, serum stability evaluation by using DLS demonstrated that the hydrodynamic size of HDMC NP remained nearly unchanged over 72 h at 37 °C, confirming its colloidal stability under physiologically relevant environments (Supporting Information Fig. S13).

The coordination structure between DOX and Fe was further investigated by comparing the Raman spectrum of HDMC NP with that of the physical mixture of UDOPA and DOX (Fig. 3G and Supporting Information Fig. S14). Based on the previously reported spectral characteristics of DOX–Fe complexes, notable changes were observed in 443/465 cm⁻¹ and 1210/1244 cm⁻¹ peak intensity ratios, indicating iron coordination of Fe to DOX. These vibrational modes have been assigned to the anthracycline chromophore of DOX, particularly to phenolic hydroxyl and carbonyl (C O) groups, rather than to the daunosamine sugar moiety. The Raman spectra of HDMC NP exhibited a lower 443/465 peak intensity ratio and a dominant peak shift from 1210 to 1244 cm⁻¹ compared with the spectra of the physical mixture of UDOPA and DOX, confirming the formation of a DOX–Fe coordination complex within the HDMC NP⁸¹.

The coordination structure of DOPA–Fe–DOX within the HDMC NP was verified by XPS (Fig. 3H). The XPS spectrum of HDMC NP revealed the Fe 2p₃ peak, similar to that observed in the XPS spectrum of UDFe. Deconvolution of the Fe 2p₃ region confirmed Fe³⁺ states, showing Fe³⁺ 2p₃/₂ and 2p₁/₂ peaks together with their corresponding satellite peaks, consistent with those observed for UDFe (Fig. 3I)^47,82,83. The XPS spectrum of HDMC NP exhibited an additional Fe³⁺–O peak, indicating a stronger coordination environment within HDMC NP than within UDFe⁸⁴. The analysis of the O 1s spectra peak further confirmed the enhanced intensity of Fe–O signals, supporting the formation of an Fe coordination complex (Fig. 3J). The presence of Fe³⁺ ions was confirmed using energy-dispersive X-ray spectroscopy (EDS) and X-ray fluorescence (XRF). As expected, no Fe signals were observed in the XRF and EDS spectra of UFH and UDOPA, whereas peaks corresponding to Fe were observed in the spectra of UDFe and HDMC NP, confirming the presence of Fe³⁺ ions (Supporting Information Fig. S15). UV–Vis spectroscopy was performed to characterize DOX, UDFe, and HDMC NP. Compared with free DOX, HDMC NP exhibited a red-shifted absorption band centered around ∼600 nm, consistent with previously reported anthracycline–Fe coordination involving the aromatic core of DOX^85,86. Such long-wavelength absorption is attributed to electronic structure changes of the anthracycline chromophore upon metal coordination rather than to the sugar moiety, confirming the formation of the coordinated HDMC nanoplatform (Fig. 3K). The anticoagulant activity of heparin was found to be abolished after DOPA conjugation and Fe³⁺ coordination, confirming that HDMC NP no longer retains the anticoagulant property of native heparin (Supporting Information Fig. S16).

3.6. Acid-triggered disruption of coordination and release of DOX release

The pH-responsive behavior of HDMC NP was evaluated by lowering the environmental pH to a mildly acidic range and analyzing structural changes. UV–Vis spectroscopy showed a decrease in absorbance at 600 nm, corresponding to the release of DOX from the DOPA–Fe–DOX coordination within HDMC NP under acidic conditions (Fig. 3L). Under acidic conditions, a visible color change was observed, with the HDMC NP exhibiting a color similar to that of free DOX, confirming the progressive release of DOX over time as supported by the reduction in absorbance at 600 nm (Fig. 3M). Consistent with the UV–Vis results, quantitative DOX release was further evaluated using a dialysis method (MWCO 3.5 kDa) at 37 °C, followed by high-performance liquid chromatography (HPLC). Time-dependent HPLC chromatograms showed a progressive increase in DOX peak intensity under acidic conditions (Supporting Information Fig. S17A), and integration of the peak areas revealed a cumulative DOX release profile that closely matched the UV–Vis-derived release trend (Fig. S17B). The DLS analysis of HDMC NP revealed that the release of DOX reduced the hydrophobic driving force for self-assembly, leading to disruption of the nanostructure and a marked increase in particle size to 1392.3 ± 620.3 nm (PDI = 1), indicating loss of nanoparticle integrity under acidic conditions (Fig. 3N).

Notably, this pH-responsive behavior further supports the DOPA–Fe–DOX coordination structure, since a control mixture prepared identically but with UFH replacing UDOPA (UFH/Fe/DOX) showed no comparable dissociation between acidic and neutral conditions (Supporting Information Fig. S18). This difference underscores the essential role of the catechol moiety in DOPA in facilitating dynamic, pH-sensitive catechol–metal–drug coordination (DOPA–Fe–DOX), which clearly differentiates HDMC NP from non-catechol-based DOX–Fe assemblies. In contrast, HDMC NPs exhibited negligible sensitivity to glutathione (GSH), as exposure to varying GSH concentrations (0–10 mmol/L) did not induce significant changes in DOX release over 24 h, indicating that coordination dissociation is primarily governed by pH rather than reductive conditions (Supporting Information Fig. S19).

3.7. Cytotoxicity profile and pathway-dependent cell death of HDMC NP

The cytotoxic effects of HDMC NP were evaluated in B16F10.OVA melanoma cells using a 48 h cell viability assay. As expected, UFH exhibited negligible cytotoxicity, whereas HDMC NP exhibited the highest cytotoxicity among all treatment groups (Fig. 4A). Notably, HDMC NP induced higher cytotoxicity than an equivalent dose of free DOX, suggesting an enhanced antitumor effect arising from the coordinated formulation (Supporting Information Fig. S20). In addition, combination index (CI) analysis performed using CompuSyn software revealed synergistic cytotoxic effects between UDFe and DOX in the HDMC NP formulation across multiple effect levels (ED₅₀–ED₉₅), confirming a synergistic interaction rather than a simple additive effect (Fig. S20). In contrast to tumor cells, HDMC NP exhibited substantially reduced cytotoxicity toward normal fibroblast cells under the same experimental conditions, indicating a favorable tumor-selective cytotoxic profile (Supporting Information Fig. S21 and Fig. S22).

Figure 4.

In vitro evaluation of the therapeutic effect of HDMC NP. (A) Cell viability of B16F10.OVA melanoma cells treated with UFH, UDOPA, UDFe, or HDMC NP for 48 h at various concentrations (n = 6). (B) Apoptosis analysis of B16F10.OVA melanoma cells treated with DOX or HDMC NP in the presence of pathway-specific inhibitors. (C) Results of Western blotting of GPX4 and SLC7A11 protein expression in B16F10.OVA melanoma cells. Quantification of (D) lipid peroxidation (BODIPY) and (E) CRT expression on the cell surface in B16F10.OVA melanoma cells, as determined by flow cytometry (n = 3). (F) Immunofluorescence images of B16F10.OVA melanoma cells showing intracellular DOX, Fe²⁺ (FerroOrange), ROS (DCF–DA), ferroptosis markers (SLC11A7 and GPX4), and the ICD marker (CRT) after treatment with control, DOX, UDFe, or HDMC NP. Scale bar = 50 μm. (G) Quantification of phagocytosis by RAW264.7 macrophages labeled with CellTracker DeepRed after co-culture with pretreated B16F10.OVA melanoma cells, as determined by flow cytometry (n = 3). (H) Representative fluorescence images of RAW264.7 macrophages (red) engulfing pretreated B16F10.OVA melanoma cells (green). Scale bar = 20 μm. (I) Flow cytometric analysis of RAW264.7 macrophage migration in a Transwell assay (n = 3). (J) Representative Hematoxylin and Eosin (H&E)-stained images of migrated RAW264.7 cells in the Transwell membrane assay. Scale bar = 200 μm. Fluorescence images were acquired at 20 × magnification using a fluorescence microscope. Channels correspond to emission at wavelengths of 457 (blue), 488 (green), and 500 nm (red). The acquisition settings were kept constant. Data are presented as mean ± SD. Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001 vs. control. ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. DOX.

To delineate the dominant cell death pathways underlying the enhanced cytotoxicity of HDMC NP, apoptosis levels were quantified using pathway-specific inhibitors. In the DOX-treated group, apoptosis levels in the free DOX-treated group (P.C), were 79.31 ± 1.29 (Fig. 4B). Relative to this value, treatment with the iron chelator deferoxamine (DFO) resulted in only a 2.98% reduction, and ferrostatin-1 (Fer-1) caused a 21.20% reduction, whereas the pan-caspase inhibitor Z-VAD-FMK led to a pronounced 49.28% decrease in apoptosis. These results indicate that DOX-induced cytotoxicity is primarily mediated by apoptosis, with limited dependence on iron availability or ferroptosis-related pathways. In contrast, HDMC NP-treated cells exhibited a distinct inhibitor response profile. Apoptosis levels in the HDMC NP-treated group (P.C) were 83.41 ± 1.44. Upon inhibitor treatment, apoptosis was reduced by 10.00% with DFO, 18.48% with Fer-1, and only 9.17% with Z-VAD-FMK, relative to the P.C group. The relatively greater sensitivity to iron chelation and ferroptosis inhibition, together with the limited effect of caspase inhibition, indicates that HDMC NP-induced cytotoxicity is predominantly iron-dependent and ferroptosis-associated, rather than apoptosis-driven.

3.8. Cellular evidence of ferroptosis and immunogenic cell death induced by HDMC NP

The induction of ferroptosis was investigated by Western blotting to determine the expression levels of GPX4 and SLC7A11, key regulators of ferroptosis, after treatment with HDMC NP at equivalent DOX concentrations (Fig. 4C). The expression level of GPX4 remained unchanged in cells treated with UDFe; however, it was significantly downregulated in HDMC NP-treated cells, indicating ferroptosis induction. Similarly, the expression level of SLC7A11 was reduced in both UDFe- and HDMC NP-treated cells, with a more pronounced decrease observed in HDMC NP-treated cells. To further examine ferroptosis-associated protein responses, Western blotting analysis was performed in cells treated with free DOX, a simultaneous DOX/Fe mixture, or HDMC NP, in the presence or absence of the ferroptosis inhibitor ferrostatin-1 (Fer-1) (Supporting Information Fig. S23). GPX4 expression was markedly reduced in HDMC NP-treated cells, whereas little change was observed following free DOX or DOX/Fe treatment. Importantly, co-treatment with Fer-1 partially restored GPX4 expression specifically in the HDMC NP group, supporting a ferroptosis-dependent regulation of GPX4. In contrast, SLC7A11 exhibited a more complex response pattern. While HDMC NP treatment was associated with a reduction in SLC7A11 expression, the effect of Fer-1 on SLC7A11 levels was less pronounced and varied across treatment groups, including DOX and DOX/Fe. Together, these results indicate that GPX4 downregulation serves as a robust marker of ferroptosis in HDMC NP-treated cells, whereas SLC7A11 expression appears to be influenced by additional regulatory factors beyond ferroptosis inhibition alone.

Lipid peroxidation, a hallmark of ferroptosis, was quantified in melanoma cells using BODIPY 581/591 C11 staining followed by flow cytometry. Significant lipid peroxidation was detected in DOX- and HDMC NP-treated cells, whereas only minimal levels were observed in UDOPA- and UDFe-treated cells. HDMC NP induced approximately 5.7 times more lipid peroxidation than that in the control and 1.7 times more than that in DOX (Fig. 4D). Consistent with these results, additional analysis comparing free DOX, DOX/Fe, and HDMC NP revealed that DOX and DOX/Fe induced comparable levels of lipid peroxidation, while HDMC NP triggered a significantly higher ferroptotic lipid peroxidation response (Supporting Information Fig. S24A). These findings were further supported by malondialdehyde (MDA) assays, which confirmed a significant increase in lipid peroxidation in HDMC NP-treated cells compared with that in control and DOX-treated cells (Fig. S24B). Consistent with the increased lipid peroxidation, the level of ROS in HDMC NP-treated cells increased by 5.1-fold compared with that in control-treated cells and by 1.8-fold compared with that in DOX-treated cells (Supporting Information Fig. S25), indicating substantial oxidative stress. In addition, cell-surface CRT expression—a marker of immunogenic cell death (ICD)—was significantly elevated in HDMC NP-treated cells, showing a 5.3-fold increase compared with that in control-treated cells and a 1.6-fold increase compared with that in DOX-treated cells (Fig. 4E). Further comparison among free DOX, DOX/Fe, and HDMC NP treatments revealed that CRT exposure induced by DOX and DOX/Fe was comparable, whereas HDMC NP elicited a markedly higher level of CRT surface expression (Supporting Information Fig. S26). These results indicate that HDMC NP not only enhances ferroptotic oxidative stress but also more effectively promotes ICD-associated signaling compared with free DOX or non-coordinated DOX/Fe treatment.

The mechanism underlying ferroptosis induction by HDMC NP was further investigated by quantifying intracellular iron accumulation using FerroOrange staining. Compared with control cells, DOX-, UDOPA-, and UDFe-treated cells showed modest increases in intracellular iron levels, whereas HDMC NP-treated cells exhibited a markedly higher iron accumulation, reaching 16.6 times that in control-treated cells and approximately twice that in DOX-treated cells, demonstrating efficient intracellular iron delivery (Supporting Information Fig. S27). Notably, although UDFe also contains iron, its intracellular accumulation remained limited, indicating that iron associated with UDFe is inefficiently internalized or rapidly cleared in the absence of DOX-mediated stabilization.

To further evaluate the accessibility of intracellular iron, an additional set of experiments was performed in the presence of the iron chelator deferoxamine (DFO) (Supporting Information Fig. S28). Upon DFO treatment, intracellular iron levels in DOX- and DOX/Fe-treated cells were reduced to a similar extent, indicating that iron accumulated under these conditions remains relatively chelatable. In contrast, HDMC NP-treated cells maintained substantially elevated intracellular iron levels even in the presence of DFO, despite equivalent iron dosing. These results suggest that the intracellular iron pool enriched by HDMC NP is stabilized through coordination with DOX, rendering it less accessible to extracellular chelation. Given the well-established redox activity of DOX, which can reduce Fe³⁺ to Fe²⁺, the accumulated Fe²⁺ detected by FerroOrange is most likely generated via DOX-mediated intracellular redox cycling rather than solely through endogenous iron transport or reduction pathways. While contributions from cellular iron-handling machinery cannot be completely excluded, the selective enhancement of Fe²⁺ accumulation in HDMC NP-treated cells supports a dominant role of coordination-enabled DOX–Fe redox interactions in driving ferroptosis.

The intracellular behavior of the HDMC NP was visualized, and key ferroptosis- and ICD-related markers were confirmed by performing immunocytochemistry using fluorescence microscopy (Fig. 4F). Fluorescence imaging at the intrinsic emission wavelength of DOX showed that HDMC NP-treated cells exhibited substantial nuclear localization of DOX, a pattern resembling that of free DOX. Since polymer-conjugated DOX is usually restricted to the cytoplasm^87,88, the pronounced nuclear distribution observed here indicates that DOX was released from HDMC NP largely in its intact form rather than in a conjugated state. FerroOrange staining further confirmed strong intracellular Fe²⁺ accumulation in the HDMC NP group, consistent with the previous flow cytometry results. 2′,7′-Dichlorofluorescin diacetate (DCF-DA) staining demonstrated elevated reactive oxygen species (ROS) levels following HDMC NP treatment, exceeding those observed with DOX alone, indicating enhanced oxidative stress. Additionally, fluorescence signals for SLC7A11 and GPX4 in the HDMC NP-treated cells decreased over time; these results were consistent with those of Western blotting and confirmed the downregulation of key ferroptosis suppressors. Notably, the level of CRT exposure was significantly increased in HDMC NP-treated cells, confirming the induction of ICD. Together, these findings demonstrate that HDMC NP induce substantial ferroptosis by increasing the accumulation of iron and ROS while simultaneously promoting ICD via CRT exposure, highlighting their potential as a dual ferroptosis–immunogenic cancer therapy.

3.9. Immunostimulatory effects of HDMC NP on macrophage-mediated phagocytosis and recruitment

The immunostimulatory potential of HDMC NP was evaluated by co-culture and Transwell assays using macrophages and pretreated B16F10.OVA melanoma cells. The phagocytosis assay revealed that the macrophage-mediated engulfment of tumor cells significantly increased when the tumor cells were pretreated with HDMC NP, with the macrophages engulfing approximately 2.3 and 3.4 times more tumor cells than tumor cells pretreated with PBS and DOX, respectively (Fig. 4G and Supporting Information Fig. S29). This effect likely reflects CRT-mediated immunogenic signaling, as indicated by the increased internalization of HDMC NP-pretreated tumor cells by macrophages (Fig. 4H).

A Transwell migration assay was performed to evaluate the recruitment of immune cells. The migration of macrophages toward HDMC NP-pretreated tumor cells increased by 1.6-fold compared with that toward PBS-pretreated cells and 1.2-fold compared with that toward DOX-pretreated cells (Fig. 4I and Supporting Information Fig. S30). Histological analysis of the transwell membrane revealed that significantly more macrophage-like RAW264.7 cells migrated toward HDMC NP-pretreated cells than toward other groups (Fig. 4J). These findings indicate that HDMC NP enhances the phagocytic activity of macrophages against tumor cells and facilitates their recruitment, demonstrating a strong immunostimulatory effect on innate immune responses.

3.10. Immune activation and antigen presentation in co-cultured splenocytes

The immunomodulatory effects of HDMC NP were evaluated by coculturing splenocytes from naïve mice with tumor cells pretreated with HDMC NP or control formulations for 24 h (Fig. 5A). Flow cytometry analysis revealed substantial changes in immune cell populations, with activation observed in both adaptive and innate compartments. Total T cells (CD3⁺) increased by approximately 1.2-fold in the HDMC NP group compared to control group (Fig. 5B). Within this population, helper T cells (CD4⁺) were significantly elevated by 1.6-fold in the HDMC NP group compared to control group (Fig. 5C), surpassing even the DOX group and indicating robust activation of CD4⁺ T cell-driven immune signaling. Cytotoxic T cells (CD8⁺) increased by 1.3-fold in the HDMC NP group compared with the control group (Fig. 5D), supporting the cytotoxic arm of the antitumor response activated helper T cells (CD4⁺CD25⁺) increased by 1.6-fold in the HDMC NP group compared with the control group (Fig. 5E), indicating antigen-specific stimulation and functional activation of T cells. Activated cytotoxic T cells (CD8⁺CD25⁺) increased significantly by 1.8-fold in the HDMC NP group compared with the control group (Fig. 5F), indicating increased readiness of effector T cells for tumor clearance.

Figure 5.

HDMC NP modulates immune cell composition in splenocytes ex vivo. (A) Schematic of the ex vivo assay. Mouse splenocytes were isolated and cocultured with B16F10.OVA melanoma cells pretreated with DOX, UDOPA, UDFe, and HDMC NP for 24 h. Flow cytometry analysis of immune cell subsets among splenocytes, including (B) CD4⁺ T cells (CD45⁺CD3⁺CD4⁺), (C) CD8⁺ T cells (CD45⁺CD3⁺CD8⁺), (D) total T cells (CD45⁺CD3⁺), (E) activated CD4⁺ T cells (CD45⁺CD3⁺CD4⁺CD25⁺), and (F) activated CD8⁺ T cells (CD45⁺CD3⁺CD8⁺ CD25⁺); (G, H) NK cells (CD45⁺NK1.1⁺) and NKT cells (CD45⁺NK1.1⁺ CD3⁺); (I) dendritic cells (CD45⁺CD11c⁺); (J) antigen-presenting dendritic cells (CD45⁺CD11c⁺SIINFEKL⁺); (K) activated dendritic cells (CD45⁺CD11c⁺CD86⁺); (L) macrophages (CD45⁺CD11b⁺f4/80⁺); (M) antigen-presenting macrophages (CD45⁺CD11b⁺f4/80⁺ SIINFEKL⁺); and (N) activated macrophages (CD45⁺CD11b⁺f4/80⁺CD86⁺). (O) UMAP analysis of splenocytes across treatment groups, illustrating population distribution patterns. Data are presented as mean ± SD (n = 3). Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001 vs. control. ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. DOX.

In the innate compartment, natural killer (NK) cells (NK1.1⁺) increased by 1.8-fold after HDMC NP exposure (Fig. 5G), indicating enhanced innate cytotoxic activity. Similarly, NKT cells (CD3⁺NK1.1⁺) increased by 1.7-fold in the HDMC NP group compared with the control (Fig. 5H), indicating that HDMC NP also stimulated cross-lineage innate-like T cell responses that may contribute to rapid tumor recognition. Dendritic cells (CD11c⁺) increased by 3.7-fold in the HDMC NP group (Fig. 5I), highlighting strong antigen-presenting cell (APC) activation. Tumor antigen-presenting dendritic cells (CD11c⁺SIINFEKL⁺) increased by 16.5-fold (Fig. 5J), reflecting potent tumor antigen capture and presentation. This was further supported by a 1.6-fold increase in activated dendritic cells (CD11c⁺CD86⁺) (Fig. 5K), which are crucial for co-stimulation and T-cell priming. Macrophages (CD11b⁺F4/80⁺) also increased by 1.6-fold (Fig. 5L), indicating enhanced recruitment or survival of phagocytic cells. The proportion of tumor antigen-presenting macrophages (CD11b⁺F4/80⁺SIINFEKL⁺) increased by 5.4-fold (Fig. 5M), highlighting the contribution of macrophages to antigen processing and immune activation. Additionally, the proportion of activated macrophages (CD11b⁺F4/80⁺CD86⁺) increased by 2.1-fold (Fig. 5N), indicating that HDMC NP promote macrophage recruitment and functional polarization toward an immunostimulatory phenotype capable of supporting T-cell activation. Finally, uniform manifold approximation and projection (UMAP) analysis of splenocytes (Fig. 5O) revealed distinct clustering patterns in the HDMC NP group, with increased populations of activated macrophages and dendritic cells and a noticeable reduction in myeloid-derived suppressor cells (MDSCs). These results demonstrate that HDMC NP reshaped the immune microenvironment to favor antigen presentation, T-cell activation, and effective innate immune engagement, highlighting their potential as a powerful immunostimulatory nanoplatform for cancer immunotherapy.

3.11. Synergistic antitumor efficacy of ferroptosis-driven metal–organic drug therapy

The in vivo antitumor efficacy of the control, DOX, UDOPA, UDFe, and HDMC NP was evaluated. B16F10.OVA tumor-bearing mice were established by subcutaneously injecting 5.0 × 10⁵ B16F10.OVA cells into the dorsal flank of C57BL/6 mice. One week after inoculation, mice were administered saline, DOX, UDOPA, UDFe, or HDMC NP, with both treatments delivering an equivalent DOX dose of 2.7 mg/kg. All formulations were administered intravenously on four occasions over a 13-day period (Fig. 6A). After 12 days, the relative tumor volume (V_t/V₀) in the control-treated mice reached 20.99 ± 9.64, with a similar result observed in UDOPA-treated mice (19.82 ± 3.93). In contrast, the relative tumor volumes in DOX- and UDFe-treated mice were reduced to 11.11 ± 7.78 and 9.22 ± 11.14. Tumor growth in the HDMC NP-treated mice was significantly suppressed, with a relative tumor volume of 2.45 ± 0.52, representing an approximately 95% reduction compared with the control group (Fig. 6B). To further assess treatment responses at the individual level, tumor growth curves of each mouse were plotted alongside body weight measurements (Fig. 6C). Whereas control, DOX, and UDOPA groups exhibited progressive tumor growth, HDMC NP treatment consistently suppressed tumor development across all mice, with minimal changes in body weight, indicating both efficacy and tolerability.

Figure 6.

In vivo antitumor efficacy and immune activation profile of HDMC NP. (A) Schematic of the in vivo antitumor treatment schedule. (B) Average relative tumor volume (V_t/V₀) of B16F10.OVA-bearing mice (n = 4). (C) Individual tumor growth curves and bodyweight of mice from each treatment group (n = 4). (D) Fluorescence images of H&E-, TUNEL-, and CD8-stained tumor sections (scale bar = 50 μm). Fluorescence images were acquired at 20 × magnification using a fluorescence microscope. Channels correspond to emission at wavelengths of 457 nm (blue) and 488 nm (green). Acquisition settings were kept constant. (E) TUNEL staining and (F) quantification of CD8⁺ staining (amount of DAPI (%)) (n = 3). (G) Level of intracellular ROS in tumor cells (n = 6). (H) Accumulation of intracellular Fe²⁺ in tumor cells (n = 6). (I) Levels of lipid peroxidation in tumor cells (n = 6). (J) Expression of CRT on the surface of tumor cells (n = 6). (K) Expression of transferrin on the surface of tumor cells (n = 6). (L) Body weight monitoring during the in vivo toxicity assessment (n = 6). (M) Pharmacokinetic profile of HDMC NP following intravenous injection (n = 3). (N–P) Quantitative biodistribution of free DOX and HDMC NP in the tumor (N), heart (O), and kidney (P) at 48 h post-injection (n = 3). All data are shown as mean ± SEM, except for (E, F), where data are shown as mean ± SD. Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001 vs. control. ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. DOX.

Cell death in the tumor tissues was histologically evaluated by hematoxylin and eosin (H&E) and terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL) staining of the tumor sections (Fig. 6D). H&E staining revealed more pronounced apoptosis of tumor tissue in HDMC NP-treated mice than in mice treated with other groups. TUNEL fluorescence staining exhibited the strongest signal in tumors of HDMC NP-treated mice, indicating a high level of apoptosis (Fig. 6E). In addition, the histological examination of the major organs, including the heart and kidney—which are known to be vulnerable to DOX-associated toxicity—as well as the liver, lung, and spleen, was performed by H&E staining. No obvious pathological abnormalities, inflammatory infiltration, or tissue damage were observed in any of these organs across all treatment groups, including free DOX and HDMC NP, indicating the absence of overt cardiotoxicity or nephrotoxicity under the tested conditions (Supporting Information Fig. S31). These results support the favorable systemic biocompatibility of HDMC NP. CD8 immunofluorescence staining demonstrated a significant increase in the infiltration of CD8⁺ cytotoxic T cells in tumors of HDMC NP-treated mice, indicating that HDMC NP induced cell death and immune cell infiltration into the TME. Quantitative analysis of CD8 staining revealed that the infiltration level of CD8⁺ T cells in tumors of HDMC NP-treated mice was 1259%, which was significantly higher than that in DOX- (218%), UDOPA- (306%), UDFe- (800%), and control-treated mice (Fig. 6F). These findings support the hypothesis that HDMC NP exerts antitumor effects via both direct cytotoxicity and immune-mediated mechanisms.

3.12. Mechanistic validation of ferroptosis and ICD induced by HDMC NP

As shown in Fig. 6G, the level of ROS in HDMC NP-treated tumor cells increased by approximately 2.1-fold compared with that in the control-treated tumor cells and by 1.6-fold compared with that in the DOX-treated tumor cells (Fig. 6G). This indicates that substantial oxidative stress, a biochemical characteristic of ferroptosis, was induced. The level of accumulated intracellular iron, as determined by FerroOrange staining (Fig. 6H), increased by 1.4-fold compared with control-treated cells and by 1.15-fold compared with DOX-treated cells, supporting the iron-dependent nature of HDMC NP-induced ferroptosis. The level of lipid peroxidation, as determined using BODIPY 581/591 dye (Fig. 6I), in HDMC NP-treated tumor cells increased by 4.7-fold compared with that in control-treated tumor cells and by 2.5-fold compared with that in DOX-treated tumor cells. These results indicate significant oxidative lipid damage, confirming ferroptosis as the dominant mechanism of cell death. Markers of ICD were also evaluated. Surface CRT, a canonical “eat me” signal recognized by APC, in the HDMC NP-treated tumor cells increased by 2.2-fold compared with that in the control-treated tumor cells and 1.3-fold compared with that in the DOX-treated tumor cells (Fig. 6J). Furthermore, the level of CD71 expression (transferrin receptor), which is often associated with active iron uptake and cellular stress, increased by 1.2-fold compared with that in control-treated tumor cells and was also higher than that in DOX-treated tumor cells (Fig. 6K), indicating increased iron metabolism and membrane remodeling under ferroptotic stress. These results indicate that HDMC NP induces ferroptosis in tumor cells through a coordinated increase in the levels of ROS, accumulated iron, and lipid peroxidation while simultaneously triggering ICD via CRT exposure and HMGB1 release. These combined effects contribute to the in vivo therapeutic efficacy of HDMC NP by increasing tumor immunogenicity and disrupting the redox balance.

3.13. Evaluation of the biosafety of HDMC NP

The in vivo safety of HDMC NP was evaluated via a biochemical toxicity study. C57BL/6 mice were intravenously injected with HDMC NP or DOX at an equivalent DOX dose of 10 mg/kg. Four days after treatment, the body weights of the mice were recorded, and blood samples were collected for further analysis. Although the body weights of DOX-treated mice continuously decreased, the body weights of HDMC NP-treated mice showed no significant changes compared with those of control-treated mice, indicating favorable biocompatibility (Fig. 6L). Blood collected from the abdominal aortic vein of DOX-treated mice on Day 4 revealed abnormally elevated levels of aspartate aminotransferase (AST), creatine kinase (CK), and lactate dehydrogenase (LDH), indicating hepatic and cardiac toxicities. In contrast, the levels of enzymes in the blood of HDMC NP-treated mice were comparable to those in control-treated mice, confirming the absence of significant liver and heart toxicity (Supporting Information Fig. S32A). Moreover, the weights of the organs of the HDMC NP-treated mice were comparable to those of the control-treated mice; however, the weights of the organs of DOX-treated mice were reduced (Fig. S32B). These results demonstrated that HDMC NP exhibited potent antitumor efficacy while minimizing systemic toxicity, indicating favorable in vivo biosafety.

In vivo system toxicity was evaluated to assess biosafety under therapeutically relevant conditions. During treatment, mice receiving free DOX or the DOX/Fe mixture showed marked body weight loss, whereas HDMC NP-treated mice maintained stable body weights comparable to those of control mice (Supporting Information Fig. S33A and S33B). Serum biochemical analysis further revealed that DOX/Fe co-treatment induced pronounced systemic toxicity, as evidenced by significant elevations in cardiac- and liver-associated biomarkers, whereas HDMC NP did not cause notable abnormalities despite equivalent dosing (Fig. S33C–S33H). These results indicate that although simple DOX/Fe co-administration can enhance cytotoxic effects, it also leads to severe systemic toxicity, highlighting the necessity of coordination-driven HDMC NP to achieve therapeutic efficacy with acceptable biosafety.

3.14. Pharmacokinetics and biodistribution of HDMC NP

The pharmacokinetics of HDMC NP were evaluated after intravenous injection at a DOX-equivalent dose of 1.35 mg/kg. Serum samples were collected at predetermined time points and analyzed based on fluorescence signals, using intrinsic DOX fluorescence for free DOX and RITC labeling for HDMC NP. Free DOX was rapidly cleared from circulation and became nearly undetectable within 30 min, whereas HDMC NP showed prolonged blood retention with detectable signals up to 72 h (Fig. 6M and Supporting Information Fig. S34).

The biodistribution of HDMC NP was further evaluated after intravenous injection at a DOX dose of 2.7 mg/kg. At 48 h post-injection, major organs and tumors were harvested, and fluorescence intensity was quantified based on DOX signals. HDMC NP exhibited significantly enhanced tumor accumulation compared with free DOX, showing approximately a 2.17-fold increase in tumor-associated fluorescence intensity (Fig. 6N). In contrast, fluorescence signals in the heart and kidney showed a trend toward higher accumulation in the free DOX group; however, these differences did not reach statistical significance (P = 0.0639 for heart and P = 0.2426 for kidney; Fig. 6O–P). Representative ex vivo fluorescence images further confirmed the distinct biodistribution profiles of HDMC NP and free DOX (Supporting Information Fig. S35A). Quantitative analysis revealed no significant difference in liver or lung accumulation between the two groups, whereas HDMC NP showed significantly reduced accumulation in the spleen compared with free DOX (Fig. S35B–S35D). Collectively, these results demonstrate that HDMC NP prolongs systemic circulation and preferentially enhances tumor accumulation of DOX without significantly increasing off-target distribution to major organs. Although heart accumulation of HDMC NP was not significantly lower than that of free DOX, the absence of cardiotoxic biomarkers and histological damage suggests that the reduced cardiotoxicity arises not solely from altered biodistribution but also from the coordination-driven formulation, which modulates intracellular iron accessibility and redox activity in non-tumor tissues. These findings indicate that both improved tumor targeting and intrinsic formulation-dependent safety contribute to the favorable therapeutic profile of HDMC NP.

3.15. Anti-tumor immune responses within the tumor microenvironment

The modulation of immune responses within the TME by HDMC NP was investigated by isolating immune cells from the tumors of treated mice and analyzing their phenotypes by flow cytometry. In tumors from HDMC NP-treated mice, the proportions of total T cells and helper T cells increased by approximately 1.3- and 1.1-fold, respectively, compared with those from control-treated mice (Fig. 7A and B). Although the increase in the level of CD4⁺ T cells was modest, it reflected a broad immune activation trend in response to HDMC NP. Substantial changes were observed in the effector T cell compartments. Activated helper T cells expressing interferon-gamma (Fig. 7C) increased by 1.5-fold, whereas CD8⁺cytotoxic T cells (Fig. 7D) increased by 2.3-fold, indicating enhanced antitumor cytotoxic activity. The levels of regulatory T cells (CD4⁺Foxp3⁺) also showed a 1.6-fold decrease, suggesting not only the induction of cytotoxicity but also an immunostimulatory reprogramming of the TME (Fig. 7E). The population of IFN-γ-positive CD8⁺ T cells significantly increased by 2.6-fold after HDMC NP treatment compared with that after control treatment (Fig. 7F). In contrast, the proportion of exhausted CD8⁺PD-1⁺ T cells in tumors from HDMC NP-treated mice decreased by approximately 20% compared with the control group, suggesting that HDMC NP may restore T-cell effector function by alleviating exhaustion (Fig. 7G). Flow cytometry revealed that the frequencies of tumor-associated dendritic cells and macrophages remained unchanged across the treatment groups (Fig. 7H–J). However, the proportion of proinflammatory M1-like macrophages expressing iNOS (Fig. 7K) increased by 1.8-fold in the HDMC NP group, whereas immunosuppressive macrophages expressing arginase-1 (Fig. 7L) decreased by ∼25% relative to the control. These shifts suggest HDMC NP reprograms the myeloid compartment toward a more inflammatory and antigen-presenting phenotype.

Figure 7.

HDMC NP promotes antitumor immune activation in the tumor microenvironment. Tumor-infiltrating immune cells were analyzed by flow cytometry after in vivo treatment with HDMC NP in B16F10.OVA tumor-bearing mice. (A, B) Percentages of total CD3⁺ T cells (CD45⁺CD3⁺) and CD4⁺ T cells (CD45⁺CD3⁺CD4⁺). (C) IFN-γ⁺ CD4⁺ T cells (CD45⁺CD3⁺CD4⁺IFN-γ⁺). (D) CD8⁺ cytotoxic T cells (CD45⁺CD3⁺CD8⁺). (E) Regulatory T cells (CD45⁺CD3⁺CD4⁺Foxp3⁺). (F) IFN-γ⁺ CD8⁺ T cells (CD45⁺CD3⁺CD8⁺IFN-γ⁺). (G) PD-1⁺CD8⁺ T cells (CD45⁺CD3⁺CD8⁺PD-1⁺). (H–J) Dendritic cells (CD45⁺CD11c⁺), activated DCs (CD45⁺CD11c⁺CD86⁺), and macrophages (CD45⁺CD11b⁺F4/80⁺). (K, L) M1 macrophages (CD45⁺CD11b⁺F4/80⁺iNOS⁺) and M2 macrophages (CD45⁺CD11b⁺F4/80⁺Arginase⁺). (M) Myeloid-derived suppressor cells (CD45⁺CD11b⁺Gr-1⁺). (N–Q) NK cells (CD45⁺NK1.1⁺), IFN-γ⁺ NK cells (CD45⁺NK1.1⁺IFN-γ⁺), NKT cells (CD45⁺NK1.1⁺CD3⁺), and IFN-γ⁺ NKT cells (CD45⁺NK1.1⁺CD3⁺IFN-γ⁺). (R) Schematic showing HDMC NP-mediated antitumor immune activation through ferroptosis-induced immunogenicity and remodeling of the TME. Data are presented as mean ± SEM (n = 8). Statistical significance was determined by one-way ANOVA with Tukey's test. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001 vs control. ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs DOX.

Myeloid-derived suppressor cells (MDSCs) (Fig. 7M) were reduced by ∼67% in the HDMC NP group, indicating a strong suppression of immunosuppressive myeloid populations. This result was accompanied by a 2.1-fold increase in NK cells (Fig. 7N) and a 2.6-fold increase in IFN-γ-producing NK cells (Fig. 7O), both of which contribute to innate cytotoxic responses. NKT cells (Fig. 7P) and IFN-γ-positive NKT cells (Fig. 7Q) increased by 2.5- and 2.0-fold, respectively. These data demonstrate that HDMC NP reshaped the immune landscape in tumors by increasing cytotoxic T and NK cell populations, restoring T cell functionality, reducing immunosuppression, and promoting innate and adaptive immune synergy, thereby contributing to a potent antitumor immune response. Representative flow cytometry dot plots corresponding to these analyses are shown in Supporting Information Fig. S36, further illustrating the distinct immune profiles across treatment groups (Supporting Information Fig. S36).

The systemic immune responses were evaluated by analyzing the immune cell profiles in the tumor-draining lymph nodes (TDLNs) (Supporting Information Fig. S37), spleen (Supporting Information Fig. S38), and peripheral blood (Supporting Information Fig. S39) of the treated mice. In all examined tissues, HDMC NP treatment consistently increased the populations of CD3⁺ T cells, CD4⁺ helper T cells, and CD8⁺ cytotoxic T cells compared to the control group, reflecting systemic T cell activation. The populations of innate immune subsets, such as NK and NKT cells, were also increased, particularly in the peripheral blood and spleen, supporting broad innate immune activation. The populations of dendritic cells and activated dendritic cells (CD80⁺) were significantly increased in the HDMC NP group across all tissues, indicating enhanced antigen-presenting capacity. In the myeloid compartment, macrophages and M1-like macrophages were enriched in both the spleen and lymph nodes, whereas immunosuppressive subsets, including MDSCs and Ly6G⁺ neutrophils, were significantly reduced. These results highlight the ability of HDMC NP to remodel both systemic and local immune landscapes and to promote effective antitumor immunity.

3.16. HDMC NP reprograms the tumor cell transcriptome and enhances immune gene signatures, as revealed by single-cell RNA sequencing

The cellular and molecular mechanisms underlying the therapeutic effects of HDMC NP were elucidated by performing single-cell RNA sequencing (scRNA-seq) on tumors isolated from treated mice. Tumor-infiltrating immune and nonimmune cells were profiled to assess the changes in gene expression, cell state, and immune context.

Differential gene expression analysis revealed that HDMC NP treatment profoundly reprogrammed the transcriptional landscape of tumor cells (Fig. 8A). Compared with the control group, HDMC NP markedly upregulated mitochondrial respiratory genes, including Nd1–Nd4, Cox1–Cox3, Cytb, and Atp6/Atp8, suggesting enhanced oxidative phosphorylation and ROS generation. Meanwhile, genes associated with ribosomal structure and transcriptional regulation (Rpl17, Rps24, Rpl35, Brd4) were downregulated, indicating global translational suppression and metabolic adaptation to oxidative stress. Together, these results demonstrate that HDMC NP reshapes tumor metabolism toward a ferroptosis-associated oxidative state. When compared with doxorubicin alone, HDMC NP induced a distinct transcriptional signature driven by the Fe³⁺–DOX redox coordination (Fig. 8B). This redox synergy activated stress-adaptive and membrane-remodeling genes (Prdm16, Npc1, Slc4a2, Tmem5, Tmem51), while repressing RNA-modifying enzymes (Mettl1, Kmt2e). These shifts reveal that Fe³⁺ coordination transforms doxorubicin from a conventional DNA-damaging chemotherapeutic into a redox-activated, ferroptosis-inducing agent that simultaneously drives metabolic and proteostatic reprogramming.

Figure 8.

Single-cell RNA sequencing reveals HDMC NP-induced transcriptomic remodeling of the TME. Single-cell RNA sequencing was performed on tumor cells isolated from B16F10.OVA-bearing mice after in vivo treatment. (A, B) Volcano plots showing differentially expressed genes between HDMC NP and control, and HDMC NP and DOX groups. (C) Gene ontology (GO) enrichment analysis highlighting significantly upregulated biological processes in HDMC NP-treated tumors. (D–F) Heatmaps showing expression profiles of genes related to ferroptosis, cell death, and immune response across control, DOX, and UDFe groups. (G) UMAP plots showing the clustering and distribution of myeloid cell populations, including M1 macrophages, M2 macrophages, monocytes, and MDSCs. (H) UMAP visualization of dendritic cell subpopulations categorized by activation markers (CD80, CD86, and CD40). (I, J) Normalized expression levels of selected marker genes across treatment groups. (I) Differential expression between control and HDMC NP-treated tumors. (J) Differential expression between HDMC NP- and DOX-treated tumors.

To further understand the functional consequences of these transcriptional changes, gene ontology enrichment analysis was performed (Fig. 8C). Upregulated genes were enriched in pathways related to oxidative stress response, lipid peroxidation, apoptotic regulation, and antigen processing and presentation, indicating that HDMC NP simultaneously activates ferroptotic and immune-modulatory signaling within tumor cells. These transcriptomic changes were further supported by heatmap analysis of representative gene sets. In the HDMC NP group, the expression levels of ferroptosis-promoting genes (Ptgs2, Acsl4, and Hmox1) were increased, whereas the expression levels of ferroptosis-resistance genes (Gpx4, Slc7a11, and Fth1) were suppressed (Fig. 8D). In addition, the expression of genes related to apoptosis and necroptosis, including Bax, Casp3, and Ripk3, increased, indicating the coactivation of multiple cell death pathways (Fig. 8E). Immune response-associated genes were strongly upregulated by HDMC NP treatment (Fig. 8F). Genes involved in interferon signaling (Stat1, Irgm1, Ifitm3), cytotoxic responses (Gzmb, Prf1), and antigen presentation via MHC class I molecules (H2-k1, H2-d1) were prominently increased, indicating enhanced immune activation and cytotoxic potential within the tumor microenvironment.

The effect of HDMC NP on immune cell populations was examined by performing UMAP-based clustering of tumor-infiltrating immune cells (Fig. 8G and H). HDMC NP group exhibited a notable increase in populations of M1-like macrophages and a reduction in populations of M2 macrophages and MDSCs, indicating the reprogramming of the myeloid compartment toward a proinflammatory phenotype. Further UMAP analysis of dendritic cells revealed increased frequencies of activated subsets expressing CD86 and CD40 in the HDMC NP group (Fig. 8H), consistent with the enhanced antigen-presenting capacity.

These immunostimulatory transcriptomic changes were further corroborated by UMAP analysis of tumor-infiltrating lymphocytes based on high-dimensional flow cytometry (Supporting Information Fig. S40). In-depth visualization of CD4⁺ T cell subsets (Fig. S40A) revealed a significant increase in the population of IFN-γ⁺CD69⁺CD4⁺ T cells and a decrease in Foxp3⁺ regulatory T cells in the HDMC NP group, indicating enhanced activation of helper T cells with reduced immunosuppressive balance. Similarly, clustering of CD8⁺ T-cell populations (Fig. S40B) revealed an increased presence of effector subsets, including CD8⁺CD69⁺IFN-γ⁺ and CD8⁺CD25⁺IFN-γ⁺ cells, together with a reduction in exhausted CD8⁺PD-1⁺ cells. In contrast, the proportion of CD8^– T cells decreased after HDMC NP treatment, indicating restoration of cytotoxic T-cell functionality. Furthermore, UMAP analysis of the innate lymphocyte compartment (Fig. S40C) demonstrated increased populations of NK and NKT cells, including IFN-γ-producing subsets, indicating the enhancement of innate effector activity.

Treatment with HDMC NP markedly altered the transcriptional program of tumor cells compared with the control group. Multiple mitochondrial respiratory genes (Nd1, Nd2, Nd3, Nd4l, Cox1, Cox2, Cox3, Cytb, Atp6, and Atp8) were significantly upregulated, indicating enhanced mitochondrial oxidative metabolism. In contrast, several ribosomal genes, including Rpl17, Rps24, Rpl21, and Rpl35, were downregulated, suggesting a suppression of global protein translation under oxidative stress conditions. Mitochondrial ribosomal components (Mt-rnr1 and Mt-rnr2) showed a modest increase, implying an adaptive response to sustain mitochondrial protein synthesis under iron-dependent metabolic stress. Furthermore, genes associated with protein folding and iron-sulfur cluster assembly (Hsp90aa1, Hspa1b, Isca1, and Fam220a) were moderately downregulated, indicating elevated proteostatic stress concurrent with intensified mitochondrial metabolism. Collectively, these findings suggest that HDMC NP treatment induces a metabolic transition characterized by enhanced oxidative phosphorylation, translational suppression, and iron-dependent stress adaptation in tumor cells (Fig. 8I).

When comparing HDMC NP with doxorubicin, distinct transcriptional profiles were observed. While Dox maintained a transcriptional pattern close to the basal level, HDMC NP induced a ferroptosis-linked transcriptional response driven by the Fe³⁺–DOX redox synergy. This coordination markedly enhanced the expression of stress-adaptive and membrane remodeling-related genes such as Ctsz, Cd44, Caprin1, Sumo2, and Strap, which are associated with oxidative stress adaptation, lysosomal remodeling, and translational regulation. Conversely, several ribosomal structural genes (Rps29, Rp7, Rpl26) and the proliferative metabolic enzyme Odc1 were downregulated, indicating a suppression of global translation and proliferation-related metabolism. Additionally, mitochondrial ATP synthase (Atp5b) and metabolic genes (Got2, Apoe) exhibited modest reduction, reflecting a shift in energy metabolism and adaptation to iron-mediated oxidative stress. In contrast, genes involved in membrane trafficking and proteostasis, such as Rpn1, Laptm4a, and Mprip, were elevated, reflecting enhanced endoplasmic reticulum and lysosomal activity for protein quality control. These findings highlight that Fe³⁺ coordination transforms DOX from a conventional chemotherapeutic into a redox-activated ferroptosis inducer, revealing the unique transcriptional signature and metabolic reprogramming conferred by the HDMC nanoplatform (Fig. 8J).

4. Conclusions

In summary, we established a supramolecular catechol–metal–drug nanoplatform (HDMC NP) in which doxorubicin (DOX) is stably coordinated with Fe³⁺ via catechol moieties on a heparin backbone, forming a coordination-defined supramolecular coordination network. In this design, the drug itself serves as a functional building block, replacing inert carriers and enabling high drug-loading efficiency. This strategy combines structural simplicity, biocompatibility, and pH-responsiveness, allowing the nanoparticles to disassemble under acidic TME conditions and release DOX in its intact molecular form.

The dual coordination between Fe³⁺ and DOX generates a redox-active catalytic core, in which sustained Fe³⁺/Fe²⁺ cycling, consistent with DOX-mediated redox activity, leads to abundant ROS generation. This synergistic redox process amplifies lipid peroxidation and ferroptotic stress far beyond what either component can achieve in the absence of defined coordination. Simultaneously, the ensuing oxidative stress is accompanied by CRT exposure and dendritic cell maturation, initiating ICD and promoting cytotoxic T-cell infiltration. Furthermore, single-cell RNA sequencing revealed that HDMC NP is associated with a shift toward ferroptosis-associated metabolic states while reshaping the tumor immune microenvironment to favor antigen presentation and effector activation.

Beyond ferroptosis induction, the catechol–metal–drug coordination motif offers modularity that can be extended to other therapeutic agents and metal ions (e.g., Cu²⁺, Zn²⁺), suggesting a potentially versatile supramolecular platform for redox-driven and immune-activating cancer therapy. Collectively, this work introduces a coordination-mediated therapeutic paradigm that couples precise materials chemistry with biological ferroptosis–immune synergy, offering a new direction for next-generation nanomedicine design.

Author contributions

Gaeun Ma and Jooho Park conceived and designed the study; Gaeun Ma, Yun-Hwa Jeong, Jae-Hyeon Lee, and Seong-Bin Yang developed the methodology; Gaeun Ma, Yun-Hwa Jeong, and Seong-Bin Yang conducted the investigation and data curation; Yeonsu Ryu and Jae-Hyeon Lee performed the formal analysis and validation; Yeonsu Ryu carried out visualization; Seong-Bin Yang developed the software; Byeong-Kook Hwang and Jae-Hyeon Lee provided resources; Gaeun Ma, Yun-Hwa Jeong, Yeonsu Ryu, Jeong Uk Choi, and Jooho Park wrote, reviewed, and edited the manuscript; Jeong Uk Choi and Jooho Park supervised the project and acquired funding; All authors discussed the results and approved the final manuscript.

Conflicts of interest

The authors declare no conflicts of interest.

Appendix A. Supporting information

The following is the Supporting Information to this article: