Cell membranes-encapsulated gadolinium-doped carbon dots for the efficient photothermal and immunotherapeutic synergistic treatment of hepatocellular carcinoma

Abstract

Hepatocellular carcinoma (HCC), a predominant subtype of liver cancer, is witnessing a rising global incidence and urgently demands the development of innovative nanoplatforms that integrate precise therapeutic and immune regulatory functions. To address the limitations of conventional monotherapies, which often suffer from inadequate tumor targeting, insufficient efficacy, and limited immune activation, this study employs a "biomimetic targeting-synergy therapy" approach. We have engineered a composite system consisting of gadolinium-doped carbon dots (Gd-CDs) enveloped with hepatocellular carcinoma cell membranes (HCM), thereby imparting homologous targeting capabilities and immune activation properties. This Gd-CDs@HCM system facilitates photothermal immunotherapy, guided by bimodal fluorescence (FL) and magnetic resonance (MR) imaging. Upon laser irradiation, Gd-CDs@HCM can induce immunogenic cell death (ICD) in tumor cells. The tumor-associated antigens (TAAs) and damage-associated molecular patterns (DAMPs) released during ICD collaboratively enhance systemic anti-tumor immunity in conjunction with HCM, achieving a primary tumor ablation rate of 84.9% and inhibiting tumor progression. Consequently, this research offers an innovative strategy for real-time monitoring and precise synergistic treatment of HCC by utilizing FL/MR bimodal imaging and integrating bionic targeting, localized thermal ablation, and immune activation functions.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s12951-026-04123-9.

Introduction

Immunotherapy for hepatocellular carcinoma (HCC) has advanced significantly in recent years, yet numerous challenges persist. Immune checkpoint inhibitors, particularly PD-1/PD-L1 inhibitors, have been integrated into second-line treatments for advanced HCC [1–3]. However, these therapies face fundamental obstacles, such as low antigen presentation efficiency and inadequate T-cell infiltration [4, 5]. Photothermal therapy (PTT)-induced immunogenic cell death (ICD) has emerged as a novel strategy to address these challenges. PTT can induce tumor cells to release tumor-associated antigens (TAAs) and damage-associated molecular patterns (DAMPs) through precise thermal ablation, thereby activating antigen-presenting cells (APCs) like dendritic cells (DCs) and initiating a systemic anti-tumor immune response [6, 7]. Based on this, photothermal immunotherapy (PTI) not only enhances the specificity of local treatments but also activates the systemic immune system to strengthen anti-tumor effects, offering a promising strategy for the radical treatment of HCC.

Despite these advancements, effective PTI for HCC faces several challenges, including the poor biodegradability of traditional noble metal nanoparticle photothermal agents and the lack of synergy between photothermal conversion efficiency and immune modulation functions [8–10]. Recently, carbon dots (CDs) have emerged as a promising alternative owing to their excellent photothermal conversion properties, biocompatibility, and ease of surface functionalization, offering a breakthrough to overcome the limitations of traditional therapies [11, 12]. CDs can serve as photothermal materials and enhance T-cell infiltration and cytotoxicity by loading immune adjuvants, facilitating photothermal immunotherapy for tumors [13]. However, most CDs are rapidly cleared by the reticuloendothelial system because of nonspecific adsorption of surface charges and lack of active targeting capabilities [14], which limits their clinical application.

Tumor cell membrane biomimetic coating technology can significantly enhance the accumulation efficiency of nanoparticles at tumor sites by mimicking the structure and function of natural biological membranes [15, 16] and utilizing homologous targeting characteristics mediated by cell membrane surface adhesion molecules [17]. Therefore, coating CDs with hepatocellular carcinoma cell membranes (HCM) can achieve homologous targeting, thus improving the tumor-targeting capability of CDs, and extending their circulation time in vivo through immune evasion characteristics mediated by membrane proteins [16, 17]. Additionally, the retention of immunogenic proteins (such as calreticulin and HSP70) and TAAs on the cell membrane provides a potential antigen reservoir for in situ vaccine effects, enhancing antigen presentation and T-cell activation, thereby reversing the tumor immunosuppressive microenvironment and further improving the precision of immunotherapy [18, 19]. At present, the tumor cell membrane biomimetic systems have been focused on single therapeutic functions, such as chemotherapy or PTT, [20, 21] which showed low immune response rates and a tendency for residual tumor cells to relapse. By integrating imaging monitoring capabilities, local thermal ablation effects, and systemic immune modulation functions, a synergistic "local-systemic" treatment model can be realized. This model combines imaging monitoring, local thermal ablation, and immune activation, achieving radical tumor treatment and overcoming the limitations of single therapies, thereby opening up an innovative avenue for PTI in cancer treatment.

Based on the inherent fluorescence (FL) and magnetic resonance (MR) imaging capabilities of gadolinium-doped carbon dots (Gd-CDs), which play a significant role in the diagnosis and real-time monitoring of nanoparticle distribution in HCC [22–24], this study proposes to coat HCM onto the surface of Gd-CDs to enable FL/MR bimodal imaging and PTI for HCC. Specifically, gadolinium diethylenetriamine pentaacetic acid (Gd-DTPA), indocyanine green (ICG), polyethylene glycol (PEG), and citric acid (CA) were utilized to synthesize Gd-CDs via a one-step microwave-assisted thermal method. The synthesized Gd-CDs not only possess excellent biocompatibility, but also have multiple functions, such as excellent photothermal performance and bimodal imaging performance [25]. Subsequently, a simple extrusion method was employed to coat HCM onto the surface of Gd-CDs, resulting in Gd-CDs@HCM. The coating of HCM endows the system with homologous targeting capabilities, significantly enhancing tumor accumulation efficiency. Meanwhile, the photothermal conversion characteristics imparted by Gd-CDs can directly ablate tumor cells and induce ICD, releasing TAAs and activating DCs, thereby enhancing the anti-tumor immune response and improving the diagnostic and therapeutic capabilities for HCC (Scheme 1). This study introduces a novel paradigm for PTI in HCC and establishes a theoretical framework alongside technical guidance for advancing tumor biomimetic nanomedicine.

Scheme 1.

The schematic illustrates the synthesis of Gd-CDs@HCM and the photothermal effect induced by Gd-CDs@HCM under 808 nm laser irradiation, which synergistically stimulates antitumor immune responses in conjunction with HCM, thereby enhancing immunotherapy

Results and discussions

Structure, photothermal conversion performance, and biosafety of Gd-CDs@HCM

In this study, a multifunctional biomimetic nanodiagnostic and therapeutic platform was developed utilizing ICG, CA, Gd-DTPA, and PEG to synthesize red-light-emitting Gd-CDs by a one-pot microwave-assisted method. Transmission electron microscopy (TEM) image (Fig. 1a) revealed that the Gd-CDs exhibited a uniformly dispersed, quasi-spherical morphology with an average particle size of 12.24 ± 1.26 nm. High-resolution TEM (HRTEM) analysis (inset of Fig. 1a) indicated a lattice spacing of 0.28 nm, corresponding to the (100) plane of graphite [26]. The X-ray diffraction (XRD) pattern (Fig. S1) displayed a characteristic peak at 24.82°, indicative of the presence of graphitic (002) planes [27]. Fourier-transform infrared (FT-IR) spectroscopy (Fig. 1b) demonstrated that the Gd-CDs retained surface functional groups from ICG, including the sulfonate group (–SO₃⁻) (peaks at 1094 cm⁻¹) [28] and PEG’s C–O–C bonds (1492 cm⁻¹) [29, 30]. Additionally, coordination between Gd³⁺ ions and DTPA caused a shift of the C = O vibration peak to 1552 cm⁻¹, confirming Gd doping [22]. X-ray photoelectron spectroscopy (XPS) analysis (Fig. 1c) confirmed the presence of C, N, O, S and Gd elements in the Gd-CDs. The high-resolution Gd 4d spectrum (Fig. 1d) exhibited two characteristic binding energies at 146.9 and 141.7 eV, corresponding to Gd 4d_3/2 and Gd 4d_5/2 states, respectively, indicating that Gd exists predominantly as Gd³⁺ within the nanostructure [31]. High-resolution XPS analysis revealed characteristic peaks in the C 1 s spectrum at 284.8 and 283.3 eV, assigned to C–O/C = C and C–C/C–H bonds, respectively. For the N 1 s spectrum, peaks located at 398.8 and 401.2 eV were attributed to pyridinic and pyrrolic nitrogen, respectively. Similarly, in the O 1 s spectrum, peaks at 530.1 and 531.6 eV corresponded to hydroxyl and carbonyl groups, respectively (Fig. e–g) [32]. The high-resolution XPS spectrum of S 2p (Fig. 1h) exhibits a pair of spin–orbit coupling at binding energies of 168.47 eV (S 2p₁/₂) and 167.27 eV (S 2p₃/₂), respectively. The positions of these binding energies are consistent with the characteristic peaks of the –SO₃⁻ functional group, indicating that sulfur in Gd-CDs predominantly exists in the form of –SO₃⁻. [28] Collectively, these findings confirm the doping of Gd³⁺, endowing Gd-CDs with MR imaging capabilities. The surface retention of hydrophilic –SO₃⁻ groups enhance water solubility, while PEG’s C–O–C bonds improve biocompatibility, laying the foundation for the application of a multifunctional biomimetic nanoplatform.

Fig. 1.

a) TEM and HRTEM images of Gd-CDs. b) FT-IR spectra of the indicated samples. c) XPS full spectrum of Gd-CDs. d–h) High-resolution spectra of various elements in Gd-CDs. i) TEM image of Gd-CDs@HCM. j) Zeta potential measurements of HCM, Gd-CDs, and Gd-CDs@HCM. k) SDS-PAGE analysis of samples (1: Gd-CDs, 2: HCM, 3: Gd-CDs@HCM). l) UV–Vis spectra of Gd-CDs and Gd-CDs@HCM. m) Emission spectra of Gd-CDs and Gd-CDs@HCM (excitation wavelength = 660 nm). n) Relaxation rate comparison between Gd-CDs@HCM and gadodiamide. o) T1-weighted MR images of Gd-CDs@HCM and gadodiamide

To establish a biomimetic targeted nanoplatform, Gd-CDs were encapsulated with HCM to form Gd-CDs@HCM. Biological TEM image (Fig. 1i) showed that Gd-CDs@HCM exhibita spherical morphology with an increased average particle size of approximately 42 nm compared with Gd-CDs. Zeta potential measurements (Fig. 1j) indicate that the surface potential of Gd-CDs is + 7.63 mV. After encapsulation with HCM, the electrostatic interaction between HCM and Gd-CDs results in a reduced surface potential of Gd-CDs@HCM (+ 3.57 mV) compared with that of Gd-CDs. SDS-PAGE analysis (Fig. 1k) revealed protein band patterns consistent with the HCM, suggesting successful membrane coating and preservation of membrane protein functionalities. UV–Vis absorption spectra (Fig. 1l) of both Gd-CDs and Gd-CDs@HCM displayed strong absorption peaks between 750–810 nm, attributable to π-π* transitions within their conjugated structures [33]. The emission spectra demonstrated that under an excitation wavelength of 660 nm, the emission wavelength for Gd-CDs and Gd-CDs@HCM is 706 nm (Fig. 1m and Fig. S2). It belongs to near-infrared emission and exhibits strong tissue penetration, low scattering, and minimal background FL interference, significantly enhancing the signal-to-noise ratio for in vivo imaging [34]. Based on these characteristics, the robust emission of Gd-CDs and Gd-CDs@HCM at 706 nm demonstrates considerable potential for deep tissue imaging. To evaluate the performance of MR imaging, the Gd³⁺ content in Gd-CDs@HCM was accurately determined using inductively coupled plasma mass spectrometry (ICP-MS) (Table S1). Subsequently, the results of MR imaging and relaxation rate (r₁) analysis (Fig. 1n, o) demonstrated that with an increase in Gd³⁺ concentration, the MR signal intensity of Gd-CDs@HCM, as well as that of the clinical contrast agent gadodiamide, was correspondingly enhanced. The longitudinal r₁ of Gd-CDs@HCM was measured at 8.87 mM⁻¹·s⁻¹, which is significantly higher than that of the clinically used gadodiamide (3.02 mM⁻¹·s⁻¹). In clinical T1-weighted MR imaging sequences, a higher r₁ value of the contrast agent correlates with increased signal intensity in the images, thereby further corroborating the potential of Gd-CDs@HCM as a bimodal imaging probe [35].

To assess the photothermal properties, the temperature elevation and stability tests were performed under 808 nm laser irradiation (1 W/cm²). A solution of Gd-CDs at 400 µg/mL rapidly increased to 64.7 ℃ within 5 min (ΔT = 35 ℃), markedly surpassing pure water (ΔT = 1.5 ℃) (Fig. 2a, c). The photothermal heating effect was positively correlated with Gd-CDs concentration and laser power density (Fig. 2a, b). The photothermal conversion efficiency (ƞ) was calculated to be 46.7% (Fig. 2d), and the temperature elevation remained stable over five on/off laser cycles (Fig. 2e). Under the irradiation of an 808 nm laser (1 W/cm²), there was no significant difference in the temperature increase of the 400 µg/mL Gd-CDs solution and the Gd-CDs@HCM solution (Fig. 2f), indicating that HCM encapsulation does not significantly impair photothermal conversion [36]. These results demonstrate that Gd-CDs@HCM retain high photothermal efficiency and show enhanced tumor targeting capabilities originated from HCM modification, providing a robust foundation for multifunctional theranostic applications [19].

Fig. 2.

a) Temperature rise curve and c) thermal images of varying concentrations of Gd-CDs subjected to 808 nm laser irradiation (1 W/cm², 5 min). b) Temperature rise curves of Gd-CDs irradiated with an 808 nm laser at different power levels, using a concentration of 200 μg/mL. d) Calculation of the photothermal conversion efficiency of Gd-CDs. e) Photothermal stability of Gd-CDs at 400 μg/mL after five cycles of laser irradiation (1 W/cm²). f) Temperature rise curves of water, Gd-CDs, and Gd-CDs@HCM under 808 nm laser irradiation (1 W/cm², 5 min). g) Comparison of cell survival rates for THLE-2 and HepG2 cells incubated with various concentrations of Gd-CDs solution over 24 h. h) Images of solutions containing different concentrations of Gd-CDs@HCM and TX-100. i) Histogram of hemolysis rates for solutions containing different concentrations of Gd-CDs@HCM. j) Alterations of BALB/c mice body weight throughout treatments. k) Pathological sections of the heart, liver, spleen, lung, and kidney from BALB/c mice treated with Gd-CDs@HCM solution over different durations

Biological safety is a critical consideration in biological experiments. Accordingly, the biosafety of Gd-CDs and Gd-CDs@HCM was assessed through cytotoxicity assays, hemolysis tests, pathological examinations, and serological analyses. To begin, the in vitro cytotoxicity of Gd-CDs was evaluated using the Cell Counting Kit-8 (CCK-8) assay in HepG2 and THLE-2 cells (Fig. 2g). Results indicated that cell viability remained above 80% even at a concentration of 200 µg/mL (p > 0.05), confirming the excellent biocompatibility of Gd-CDs. The blood compatibility of Gd-CDs@HCM was subsequently evaluated through an in vitro hemolysis assay (Fig. 2h, i). Even at a concentration of 400 µg/mL, the hemolysis rate remained below 5%, demonstrating favorable blood compatibility. To validate systemic safety, six- to eight-week-old BALB/c mice were used as in vivo models. Body weight monitoring over 28 d (Fig. 2j) revealed no significant differences between treated groups and controls (p > 0.05), indicating that neither Gd-CDs nor Gd-CDs@HCM caused systemic metabolic disturbances. Histological analysis of major organs, including the heart, liver, spleen, lungs, and kidneys, using hematoxylin and eosin (H&E) staining, showed intact tissue architecture and normal cellular morphology, with no evidence of inflammatory infiltration or tissue damage (Fig. 2k and Fig. S3). Furthermore, serum biochemical analyses of hematological parameters (RBC, WBC, PLT, HGB), liver function markers (ALT, AST), and renal function indicators (UREA, CREA) (Fig. S4) revealed no statistically significant differences compared with controls (p > 0.05). All values remained within normal reference ranges, excluding the risk of hepatotoxicity and nephrotoxicity [37]. In summary, these findings collectively demonstrate that both Gd-CDs and Gd-CDs@HCM exhibit excellent biosafety both in vitro and in vivo settings.

Bimodal imaging of Gd-CDs@HCM

FL imaging and HCC cell targeting capability of Gd-CDs@HCM was investigated at the cellular level. As demonstrated in Fig. 3a, both Gd-CDs and Gd-CDs@HCM exhibited FL upon excitation at 647 nm. However, the FL intensity of Gd-CDs@HCM was significantly higher, suggesting enhanced cellular uptake in HepG2 cells compared with Gd-CDs, demonstrating the effective FL imaging capability of Gd-CDs@HCM. Furthermore, as illustrated in Fig. S5, HCC cells (Hepa1-6) showed markedly increased uptake of Gd-CDs@HCM compared with normal liver cells (THLE-2), indicating its potential for selective targeting of Hepa1-6 cells. The in vivo distribution and metabolism of Gd-CDs@HCM were further examined using small-animal FL imaging (Fig. 3b and Fig. S6). After intravenous injection for 6 h, the FL intensity at the tumor site significantly increased, reaching a peak at 12 h. This confirms that Gd-CDs@HCM can accumulate in HCC tissues through the enhanced permeability and retention (EPR) effect and the active targeting capability of HCM [38]. This enrichment of Gd-CDs@HCM in liver tumor tissues was further supported by evidence of FL signal localization. Simultaneously, FL signals in the liver and kidneys reached their metabolic peaks, while weak signals in the lungs suggested partial traversal of the blood-air barrier. FL signals in the liver and kidneys gradually diminished over time, and the whole-body FL signals were nearly undetectable after 24 h post-injection, indicating efficient clearance via hepatic metabolism and renal filtration [39]. These findings demonstrate the favorable biosafety of Gd-CDs@HCM.

Fig. 3.

a) Bright field, FL, and merge images of HepG2 cells treated with Gd-CDs and Gd-CDs@HCM. b) Time-dependent in vivo FL imaging of HepG2 tumor-bearing mice post-treatment with Gd-CDs@HCM. c) MR images of HepG2 cells treated with different concentration of Gd-CDs@HCM. d) In vivo MR imaging of HepG2 tumor-bearing mice after intravenous injection of Gd-CDs@HCM, captured at 0 and 6 h post-administration

The MR imaging capability of Gd-CDs@HCM was validated at both in vitro and in vivo levels. As shown in Fig. 3c, the MR signal intensity of Gd-CDs@HCM exhibited a positive correlation with its concentration. Subcutaneous tumors in HepG2 tumor-bearing mice displayed well-defined margins and significant T1 signal enhancement (Fig. 3d) after post-injection for 6 h, confirming its ability to target HCC lesions and achieve high-contrast MR imaging. In general, Gd-CDs@HCM exhibited excellent bimodality imaging performance in both in vitro and in vivo systems. FL imaging provided high sensitivity for real-time, dynamic tracking of tumor metabolic activity, while MR imaging offered high spatial resolution for precise localization of deep lesions. Thus, the FL/MR bimodal imaging capability of Gd-CDs@HCM can be utilized to validate its biomimetic targeted effect on HCC, providing a potential strategy for the future implementation of imaging-guided diagnosis and treatment of deep HCC.

Antitumor activity of Gd-CDs@HCM in vitro

To assess the antitumor efficacy of Gd-CDs@HCM, this study employed a comprehensive analysis involving cytotoxicity assays, live/dead cell staining, apoptosis evaluation, and migration and invasion inhibition experiments. Initially, the cytotoxicity assay using the CCK-8 method (Fig. 4a) revealed that, the inhibitory effects of various treatments on THLE-2 cells were relatively mild compared with HepG2 cells. In both the control group and the Gd-CDs (400 μg/mL) group, the survival rate of HepG2 cells exceeded 80%. In contrast, viability decreased to 63.7% in the Laser group, which is primarily attributed to direct biological damage resulting from the absorption of laser energy [40]. However, the combination of Gd-CDs, Gd-CDs@HCM, and laser irradiation resulted in a substantial increase in apoptosis of HepG2 cells, particularly in the Gd-CDs@HCM + Laser group, where the apoptosis rate reached 98.6%, indicating a significant cytotoxic effect. This finding suggests that precise PTI can be achieved by leveraging Gd-CDs@HCM’s ability to target HCC tissues, and combining with laser. To further illustrate the antitumor effects of Gd-CDs@HCM, Calcein AM/PI live/dead cell dual staining was conducted (Fig. 4b, c). Results showed a higher proportion of viable cells in the control, Gd-CDs, and Gd-CDs@HCM groups, while nearly complete apoptosis (PI positivity rate: 94.4%) was observed in the Gd-CDs@HCM + Laser group, significantly exceeding that of the single treatment groups (p < 0.01) and confirming the potent antitumor capability of Gd-CDs@HCM-mediated PTI. This enhanced effect is attributed to the following synergistic mechanisms: the targeting ability of Gd-CDs@HCM promotes its accumulation within tumor cells; laser irradiation triggers a strong photothermal effect, directly killing tumor cells; simultaneously, the photothermal effect induces ICD, further amplifying the photothermal‑immunotherapy synergy. Therefore, the observed enhancement of PI signal and cell death in the Gd‑CDs@HCM + Laser group results from the combined action of PTT and immune activation. To further elucidate the mechanism of cell death, apoptosis was quantified using Annexin V/PI flow cytometry (FCM) (Fig. 4d and Fig. S7). The findings indicated low apoptosis rates in the control, Gd-CDs, and Gd-CDs@HCM groups. Conversely, the combination of Gd-CDs, Gd-CDs@HCM, and laser irradiation led to a notable increase in apoptosis, with the Gd-CDs@HCM + Laser group showing the highest apoptosis rate, significantly exceeding that of the single‑treatment groups (p < 0.01). In the Gd-CDs@HCM + Laser group, late apoptotic cells accounted for 72.9%, while early apoptotic cells represented only 1.0%. This suggests that Gd-CDs@HCM-mediated PTI can rapidly induce apoptosis in tumor cells, thereby achieving effective tumor cell eradication. The data imply that while Gd-CDs@HCM alone exhibits limited cytotoxicity against tumor cells, and its combination with photothermal effects significantly enhances the targeted killing efficacy of immunotherapy.

Fig. 4.

a) Comparison of cell survival rates after co incubation with THLE-2 and HepG2 cells in different treatment groups for 24 h. b, c) FL intensity histograms and FL images of Calcein AM and PI staining in different treatment groups of HepG2 cells. d) The apoptosis rate of HepG2 cells co cultured with different treatment groups. e, f) The effect of different treatment groups on the invasion of HepG2 cells. *p < 0.05, **p < 0.01

To evaluate the ability of Gd-CDs@HCM-mediated PTI to inhibit the migration and invasion of HCC cells, scratch and Transwell invasion assays were performed. The scratch assay results (Fig. S8, S9) revealed that the change in scratch width in the Gd-CDs group was comparable to that of the control group, indicating that Gd-CDs alone do not impede cell migration. In contrast, the other experimental groups displayed varying degrees of delayed scratch width changes after 24 h, accompanied by reduced cell migration distances, suggesting their ability to suppress HepG2 cell migration. Among these, the Gd-CDs@HCM + Laser group exhibited the most pronounced effect (p < 0.01), demonstrating the strongest inhibition of HepG2 cell migration. This finding suggests that the photothermal effect may interfere with signaling pathways associated with cell migration within the tumor microenvironment [41]. The Transwell invasion assay results (Fig. 4e, f) further supported these observations. The number of invasive cells in the Gd-CDs@HCM + Laser group decreased by 95.0% compared with the control group, a significantly greater reduction than observed in the single treatment groups (p < 0.01). Single laser irradiation may indirectly affect cell migration capacity by inducing cellular stress and disrupting cytoskeletal architecture [40]. These findings indicate that Gd-CDs@HCM-mediated PTI effectively enhances the suppression of tumor cell migration and invasion. In summary, laser irradiation serves as a potent trigger for the synergistic antitumor effects of Gd-CDs@HCM, significantly inducing tumor cell death while concurrently inhibiting cell migration and invasion.

Gd-CDS@HCM-mediated PTI activated in vitro antitumor immunity

To investigate the activation of anti-tumor immunity mediated by Gd-CDs@HCM-induced PTI in vitro, HepG2 cells were co-cultured with Gd-CDs@HCM and subsequently exposed to an 808 nm laser (1 W/cm²), and the control group without any treatment.HepG2 cell from both the control and Gd-CDs@HCM + Laser groups were collected and subjected to RNA sequencing to analyze their gene transcriptomes. This analysis aimed to explore changes in the expression of immune-related genes. The volcano plot analysis of differential gene expression (Fig. 5a) revealed that, compared with the control group, 386 genes were upregulated and 81 genes were downregulated in the Gd-CDs@HCM + Laser group, indicating significant transcriptional alterations in tumor cells following PTI treatment. The heatmap of differentially expressed genes (DEGs) (Fig. 5b) showed that genes such as IRF1, NFKB1, and CXCL16 were upregulated in the Gd-CDs@HCM + Laser group compared with the control group. This suggests an enhancement of interferon and chemokine signaling, which may facilitate T cell activation and infiltration. The upregulation of IL7R supports T cell survival, thereby strengthening anti-tumor immunity [43]. Conversely, the downregulation of FN1 and ANGPTL3 may reduce fibrosis and pathological angiogenesis, improving immune cell infiltration and modulating the tumor microenvironment [44]. Gene ontology (GO) enrichment analysis (Fig. 5c) indicated that the DEGs in the Gd-CDs@HCM + Laser group were enriched in pathways related to the positive regulation of tumor PTI, such as T cell differentiation, cytokine activity, chemotaxis, positive regulation of cytokine production, cellular response to heat, and apoptosis. Subsequent kyoto encyclopedia of genes and genomes (KEGG) enrichment analysis (Fig. 5d) revealed that the Gd-CDs@HCM + Laser group exhibited enrichment in the MAPK signaling pathway, cytokine-receptor interaction pathway, and Th17 differentiation pathway compared with the control group. This indicates that multiple pathways associated with stress response and immune activation were significantly upregulated in tumor cells following Gd-CDs@HCM-mediated PTI, thereby enhancing the anti-tumor immune response.

Fig. 5.

a) Volcano map illustrating gene expression differences between the control group and the Gd-CDs@HCM + Laser treated group (n = 3). b) Heat map displaying DEGs in the control group compared with the Gd-CDs@HCM + Laser treated group. c) GO enrichment analysis of DEGs in the control group versus the Gd-CDs@HCM + Laser treated group. d) KEGG pathway enrichment analysis of DEGs in the control group versus the Gd-CDs@HCM + Laser treated group

To further investigate the mechanism of action of PTI mediated by Gd-CDs@HCM, key DAMPs such as high mobility group box 1 (HMGB1), adenosine triphosphate (ATP), and calreticulin (CRT) were identified as crucial elements released during the ICD process in tumor cells [45]. Immunofluorescence and ELISA assays were subsequently performed (Fig. 6a–c and Fig. S10). The results demonstrated that, compared with the control group, treatment with either Gd-CDs or Gd-CDs@HCM alone did not induce significant changes in the release of CRT, HMGB1, or ATP. In contrast, increased release of these key ICD markers was observed in the Laser group, the Gd-CDs + Laser group, and the Gd-CDs@HCM + Laser group. Further intergroup comparisons revealed that the Gd-CDs + Laser group exhibited significantly enhanced release of ICD markers relative to the Laser group (p < 0.05), indicating that Gd-CDs, as a photothermal agent, effectively amplify the photothermal effect and promote the induction of ICD. More importantly, the Gd-CDs@HCM + Laser group showed a further significant increase in CRT exposure, as well as ATP and HMGB1 release, compared with the Gd-CDs + Laser group (p < 0.01). This provides strong evidence that Gd-CDs@HCM, owing to its superior tumor-targeting capability, achieves more efficient accumulation within tumor tissues and thereby induces ICD more robustly. Additionally, FCM analysis (Fig. S11) demonstrated that CRT exposure on the surface of tumor cells in the Gd-CDs@HCM + Laser group was 2.7 times higher than in the Gd-CDs group and 11.6 times higher than in the control group (p < 0.01). ATP released into the extracellular space functions as a "find-me" signal, attracting DCs to the tumor microenvironment and facilitating their maturation, while CRT acts as an "eat-me" signal to enhance the phagocytosis of dying cells by DCs [46]. Furthermore, HMGB1 exerts a potent pro-inflammatory effect by binding to TLR4 on the surface of DCs, thereby stimulating the effective processing and cross-presentation of tumor antigens from dying cells [47]. These findings suggest that the combination of Gd-CDs@HCM and laser irradiation can effectively induce ICD in tumor cells.

Fig. 6.

a) The laser confocal scanning microscope (CLSM) image illustrates CRT exposure in HepG2 cells and the release of HMGB1 across different treatment groups. b, c) The levels of DAMPs in the supernatant of the culture medium were analyzed using ELISA (n = 3). d) Diagram of the Transwell co-culture experiment. e) FCM results showing the maturation of DCs. f–h) Cytokine levels in the culture medium were quantified by ELISA (n = 3). Data in panels (b, c, f, g, and h) are presented as mean ± standard deviation. *p < 0.05, **p < 0.01

Given the involvement of HMGB1, CRT, and ATP in DCs maturation, the proportion of mature DCs was further analyzed to evaluate their maturation status. A Transwell co-culture chamber was utilized to assess the immunostimulatory capacity of Gd-CDs@HCM under laser irradiation, with Hepa1-6 cells placed in the upper chamber and DCs in the lower chamber (Fig. 6d). FCM analysis (Fig. 6e and Fig. S12) demonstrated a 2.9-fold upregulation of CD80⁺ and CD86⁺ expression levels in DCs from the Gd-CDs@HCM + Laser group compared with the control group, indicating a substantial increase in the percentage of CD80⁺ CD86⁺ cells. DCs, as specialized antigen-presenting cells, play a critical role in orchestrating T cell-mediated immune responses. After phagocytosing antigens, immature DCs process these antigens into peptides, mature during their transit through lymph nodes, and present the peptides to T cells. This interaction drives the differentiation of T cells into effector T cells that target and eliminate tumor cells [48]. Laser-induced ICD generates TAAs, thereby enhancing DC cross-presentation capabilities, promoting their maturation, and ultimately amplifying the immune response.

Additionally, ELISA analysis (Fig. 6f–h) revealed significantly elevated levels of IL-6, TNF-α, and IFN-γ, cytokines associated with DC activation, in the Gd-CDs@HCM + Laser group. Specifically, IL-6, TNF-α, and IFN-γ levels increased by 3.8-fold, 3.5-fold, and 1.6-fold, respectively, compared with the control group. These findings suggest that Gd-CDs@HCM-induced PTI effectively promotes DC maturation and activates anti-tumor immune responses.

Antitumor activity of Gd-CDs@HCM in vivo

To evaluate the in vivo therapeutic potential of Gd-CDs@HCM, this study utilized subcutaneous HepG2 tumor-bearing BALB/c mouse models and subcutaneous Hepa1-6 tumor-bearing C57BL/6 J mouse models to assess the anti-tumor efficacy of Gd-CDs@HCM. First, a subcutaneous HepG2 tumor model in BALB/c mice was established. As shown in Fig. 7a, when the tumor volumes reached 60–100 mm³ (defined as Day 0), Gd-CDs@HCM (5 mg/kg) was administered via tail vein injection. After post-injection for 12 h, the tumor site was irradiated with an 808 nm laser (1 W/cm², 5 min). This treatment protocol was repeated on Day 3, and the experiment concluded on Day 14. Tumor tissues were collected for H&E staining (Fig. S13), which revealed irregular tumor cell morphology, disorganized arrangement, and cellular atypia, confirming the successful establishment of the HepG2 subcutaneous tumor model in BALB/c mice. Infrared thermal imaging was employed to monitor temperature changes at the tumor site (Fig. 7b, c). The results indicate that the temperature at the tumor site in the saline group, Gd‑CDs group, Gd‑CDs@HCM group, and the Laser group exhibited only a slight increase to approximately 36 ℃. In contrast, the temperatures in both the Gd‑CDs + Laser group and the Gd‑CDs@HCM + Laser group were significantly elevated (p < 0.01), confirming the effective photothermal conversion capability of Gd‑CDs and Gd‑CDs@HCM. Further comparisons revealed that the temperature increase in the Gd‑CDs@HCM + Laser group was particularly pronounced. This can be attributed to the superior tumor-targeting capability conferred by the homologous cell membrane surface modification, which facilitates greater accumulation of nanoparticles at the tumor site. In this group, the temperature in the tumor region rapidly escalated from an initial 34.2 to 61.3 ℃. It is known that local temperatures exceeding 50 ℃can induce protein denaturation and result in coagulative necrosis of tumor tissue [49]. Thus, the photothermal effect induced by Gd-CDs@HCM caused significant thermal damage to tumor tissue. After 14 d of treatment, mice in all groups maintained stable body weights (Fig. S14), indicating the good biocompatibility of Gd-CDs@HCM. Analysis of tumor volume progression and final tumor weights (Fig. 7d–f) revealed no significant anti-tumor effects in the Gd-CDs group compared with the control. However, the Gd-CDs@HCM group exhibited a marked increase in tumor growth inhibition rate (TGI), which can be attributed to the enhanced tumor localization provided by HCM encapsulation. Additionally, residual immunomodulatory molecules on the HCM surface activated systemic anti-tumor immunity, thereby amplifying therapeutic efficacy. At the same time, the tumor suppression effects observed in the Laser group primarily stem from the biological effects induced by the absorption of laser energy, such as tumor vascular damage and disruption of the cytoskeletal structure [40, 42]. These effects collectively contribute to the mortality of a portion of the tumor cells. Notably, the Gd-CDs@HCM plus laser group achieved a TGI of 98.2%, significantly outperforming individual treatments. The combination of Gd-CDs@HCM with laser irradiation facilitated tumor ablation through hyperthermia while simultaneously promoting the release of TAAs and other immunogenic factors, thereby activating immune responses and generating an “in situ vaccine” effect [50]. These findings suggest that Gd-CDs@HCM mediates efficient targeted anti-tumor activity through PTI, offering a promising strategy for precise HCC treatment.

Fig. 7.

a) Schematic diagram of the HepG2 tumor-bearing mice treatment protocol. b) Thermographic images of mice in each groups. c) Temperature–time curve of mice in each group. d) Tumor weight of HepG2 tumor-bearing mice in each group. e) Photographs of HepG2 tumor-bearing mice in each group. f) Tumor volume change curve. g) Schematic diagram of Hepa1-6 tumor-bearing mice treatment protocol. h) Weight change curve of Hepa1-6 tumor-bearing mice in each group over 14 d. i) Tumor volume change curve. j) Tumor weight of HepG2 tumor-bearing mice in each group. k) Photographs of Hepa1-6 tumor-bearing mice in each group. l) H&E, TUNEL, and Ki67 staining images of Hepa1-6 tumor-bearing mice in each group post-treatment. (G1 Saline, G2 Gd-CDs, G3 Gd-CDs@HCM, G4 Laser, G5 Gd-CDs + Laser, G6 Gd-CDs@HCM + Laser). *p < 0.05, **p < 0.01

To validate the reliability of Gd-CDs@HCM in laser-triggered HCC therapy, its therapeutic efficacy was evaluated in Hepa1-6 tumor-bearing C57BL/6 J male mice (n = 5 per group) according to the treatment protocol illustrated in Fig. 7g. Throughout the treatment period, the body weights of the mice remained stable, as shown in Fig. 7h. On day 14 post-treatment, histopathological examinations of major organs—including the heart, liver, spleen, lungs, and kidneys (Fig. S15)—alongside complete blood counts and biochemical assays (Fig. S16), confirmed that all parameters were within normal ranges, demonstrating good biocompatibility of the treatment. Tumor progression was assessed through tumor volume curves (Fig. 7i), tumor weights (Fig. 7j), and photographic evidence of tumor morphology (Fig. 7k and Fig. S17), which collectively indicated that the Gd-CDs@HCM combined with laser irradiation significantly enhanced therapeutic outcomes compared with other treatment groups. Further analysis of tumor tissues via H&E staining, TUNEL assay, and immunohistochemical evaluation of Ki-67 expression (Fig. 7l) revealed a substantial reduction in tumor cell density in the Gd-CDs@HCM + Laser group. The treated tissues exhibited pronounced degenerative changes, including cell swelling, cytoplasmic vacuolization, increased eosinophilia, and prominent nuclear pyknosis and fragmentation. When combined with laser irradiation, both Gd-CDs and Gd-CDs@HCM showed a significant increase in TUNEL-positive apoptotic cells and a marked decrease in Ki-67 expression. Notably, the Gd-CDs@HCM + Laser group exhibited the highest tumor cell apoptosis rate and the lowest Ki-67 expression. Ki-67, a well-established indicator of cellular proliferation, further corroborated the ability of the combined therapy to effectively suppress tumor cell proliferation while promoting apoptosis. These findings collectively highlight the potent tumoricidal activity of Gd-CDs@HCM under laser irradiation, underscoring its promise as an effective therapeutic strategy for HCC.

GD-CDS@HCM-mediated PTI activated in vivo antitumor immunity

The potential of Gd-CDs@HCM to stimulate systemic anti-tumor immune responses in the Hepa1-6 tumor model was assessed using immunofluorescence staining and FCM.Tumor tissues, lymph nodes, spleens, and blood samples were collected from various treatment groups for the analysis of relevant immunological markers. From the immunofluorescence images of tumor tissues (Fig. 8a), significant enhancement of CRT FL signals was observed in the Gd-CDs@HCM + Laser group, effectively promoting the translocation of HMGB1 from the nucleus to the cytoplasm and extracellular space. The synchronized increase in CRT exposure and HMGB1 release indicates that Gd-CDs@HCM combined with laser irradiation can induce ICD in tumor cells [43]. FCM results (Fig. 8b and S18) revealed that the Gd-CDs@HCM combined with laser irradiation group demonstrated a significant increase in the proportion of mature dendritic cells (DCs; CD80⁺ CD86⁺), showing a 3.1-fold increase in the lymph nodes and a 2.4-fold increase in the spleen compared with the control group. This indicates a substantial enhancement in antigen-presenting capacity [51]. Furthermore, the proportion of CD8⁺ T cells was significantly elevated in the Gd-CDs@HCM plus laser group, suggesting that Gd-CDs@HCM-mediated PTI effectively activates T cell-mediated immune responses. This activation is attributed to laser-induced tumor cell ablation facilitated by Gd-CDs@HCM, which leads to the release of TAAs. These antigens are subsequently captured by antigen-presenting cells, primarily DCs, in the spleen and lymph nodes, promoting DC maturation and the presentation of tumor antigens to T cells, thereby initiating anti-tumor immunity [50]. Additionally, the combination of Gd-CDs@HCM and laser irradiation exerted the strongest inhibitory effect on regulatory T cell (Treg; CD4⁺ CD25⁺ Foxp3⁺) populations within the tumor microenvironment, whereas treatments with either laser or Gd-CDs@HCM alone had the minimal suppression of Tregs. Immunofluorescence staining (Fig. 8c and S19) further confirmed that the Gd-CDs@HCM + Laser group exhibited the highest infiltration of CD8⁺ T cells and the lowest proportion of Foxp3⁺ Tregs. These results demonstrate that Gd-CDs@HCM-mediated PTI not only activates T cell immune responses but also mitigates the immunosuppressive tumor microenvironment. In summary, the combination of Gd-CDs@HCM and laser irradiation in Hepa1-6 tumor-bearing mice produced a robust anti-tumor immune response characterized by increased DC maturation, enhanced infiltration of CD8⁺ T cells, and reduced Tregs populations within the tumor.

Fig. 8.

a) Immunofluorescence staining images of CRT and HMGB1 within tumor tissue. b) FCM analysis of mature DCs in lymph nodes (CD11c⁺ CD80⁺ CD86⁺) and spleen (CD11c⁺ CD80⁺ CD86⁺), CD8⁺ T cells in tumors (CD3⁺ CD8⁺), and Tregs in tumors (CD4⁺ CD25⁺ Foxp3⁺) (n = 3). c) Immunofluorescence staining images of CD8⁺ T cells and Foxp3⁺ T cells within tumor tissue. d–g) Levels of TNF-α, IFN-γ, IL-6, and IL-10 in the peripheral blood of mice subjected to different treatments. Data are presented as mean ± standard deviation. *p < 0.05, **p < 0.01

ELISA was utilized to quantify the levels of TNF-α, IFN-γ, IL-6, and IL-10 in the peripheral blood of mice (Fig. 8d–g). The findings revealed that, compared with the control group, the Gd-CDs@HCM + Laser group demonstrated a 6.1-fold increase in IFN-γ levels (p < 0.01), indicating activation of anti-tumor immune responses by PTI [43]. Pro-inflammatory cytokines TNF-α and IL-6 were elevated by 8.4-fold and 6.3-fold, respectively; the synergistic effect of IL-6 and TNF-α can enhance T cell activation. Conversely, the levels of the immunosuppressive cytokine IL-10 were reduced. Given that IL-10 is secreted by Th2 cells, Tregs, and tumor-associated macrophages, its reduction suggests that Gd-CDs@HCM-mediated PTI may inhibit Tregs activity or decrease M2 macrophage polarization, thereby disrupting the immunosuppressive tumor microenvironment [52]. In conclusion, Gd-CDs@HCM combined with laser irradiation can induce ICD in tumor cells, and it activates the antitumor immune response in vivo by upregulating pro-inflammatory cytokines, downregulating immunosuppressive factors, and promoting the maturation of DCs within immune-related organs.

Conclusion

This study develops a multifunctional biomimetic nanoplatform, HCM, coated on the surface of Gd-CDs, designed for real-time monitoring of HCC using FL/MR imaging, as well as integrated PTT and immune activation. The findings reveal that Gd-CDs@HCM emits efficient red FL and exhibits strong absorption within the 750–810 nm range, achieving a photothermal conversion efficiency of up to 46.7% under 808 nm laser irradiation. With a high longitudinal relaxation rate of 8.87 mM⁻¹ s⁻¹, this composite demonstrates excellent FL/MR imaging capabilities, providing a reliable tool for the real-time monitoring of tumor targeting and therapeutic responses. The HCM coating enhances tumor accumulation through homologous targeting and prolongs blood circulation time via immune camouflage. Additionally, the residual immune signaling molecules on the HCM surface, together with local photothermal ablation, induce the release of endogenous antigens through ICD. This process synergistically promotes the maturation of DCs, the infiltration of CD8⁺ T cells, and the suppression of Tregs, thereby reversing the tumor’s immune-suppressive microenvironment. The combination of Gd-CDs@HCM with laser irradiation significantly inhibits primary tumor growth in both HepG2 and Hepa1-6 tumor-bearing mouse models. Overall, this research establishes a PTI nanobiomimetic platform for HCC, addressing the limitations of traditional nanotherapy’s singular functionality by integrating FL/MR bimodal imaging, biomimetic targeting, local thermal ablation, and immune activation within a single platform. This approach enhances anti-tumor immunity and offers a novel strategy for the precise treatment of HCC and other solid tumors.

Materials and methods

Chemicals

Indocyanine green (ICG) and citric acid (CA) were obtained from Shanghai Aladdin Biochemical Technology Co., Ltd. Polyethylene glycol (PEG), gadolinium diethylenetriamine pentaacetic acid (Gd-DTPA), and fetal bovine serum (FBS) were sourced from Sigma Corporation, USA. Flow cytometry antibodies, including CCK-8, CD11C-FITC, CD80-PE, and CD86-APC, were procured from Elabscience Biotechnology Co., Ltd. (Wuhan, China). The Annexin V-FITC-PI Apoptosis Analysis Kit was procured from Beijing LABLEAD Inc. (Beijing, China). The enzyme-linked immunosorbent assay (ELISA) kits for IL-10, TNF-α, IL-6, and IFN-γ were purchased from Amoy Lunchangshuo Biotech Co., Ltd. (Xiamen, China). Calreticulin Rabbit Monoclonal Antibody (mAb) and HMGB1 Rabbit mAb were obtained from Zen-Bioscience Co., Ltd. (Chengdu, China). Goat Anti-Rabbit IgG H&L was obtained from Bioss Biotechnology Co., Ltd. (Beijing, China). Mouse tumor infiltrating tissue lymphocyte isolation kit was obtained from Beijing Solarbio Science & Technology Co., Ltd. (Beijing, China). Minimum Essential Medium (MEM), Dulbecco’s Modified Eagle Medium (DMEM) were acquired from Zhong Qiao Xin Zhou Biotechnology Co., Ltd. (Shanghai, China). Phosphate-buffered saline (PBS), human hepatoma cells (HepG2), mouse hepatoma cells (Hepa1-6), and normal human liver cells (THLE-2) were acquired from Procell Co., Ltd. (Wuhan, China). BALB/c and C57BL/6 J mice were purchased from Beijing VitalRiver Experimental Animal Technology Co., Ltd. (Beijing, China).

Preparation of Gd-CDs

A mixture of 5 mg of ICG, 0.2 g of CA, 50 mg of PEG, and 30 mg of Gd-DTPA was added to 15 mL of deionized water. The solution underwent ultrasonic treatment for 40 min, followed by microwave heating for 3 min, obtaining a transparent deep green solution. The crude product was filtered using a disposable filter membrane with a pore size of 0.22 μm and then dialyzed. The dialyzed product was freeze-dried to obtain Gd-CDs powder.

Preparation of Gd-CDs@HCM

A membrane protein extraction reagent containing phenylmethylsulfonyl fluoride (PMSF) was added to HepG2 cells, which were thoroughly resuspended and placed on ice for 15 min. The cell suspension was transferred to a pre-chilled glass homogenizer and homogenized 30 times. The lysate was subjected to differential centrifugation at 4 ℃ to collect the final precipitate, which was resuspended in physiological saline to obtain the HCM suspension. The HCM and Gd-CDs powder were mixed in a mass ratio of 1:2. Using an Avanti mini extruder, the mixture was extruded 20 times through a 400 nm polycarbonate (PC) membrane, followed by sequential extrusion through 100 and 50 nm PC membranes, resulting in the preparation of Gd-CDs@HCM.

Characterization of Gd-CDs and Gd-CDs@HCM

Transmission electron microscopy (TEM) analysis was performed using JEM-2010 and HITACHI HT7800 electron microscopes. The zeta potential of the samples was determined via dynamic light scattering (Nano-ZS90, Malvern, UK). Fourier-transform infrared spectroscopy (FT-IR; TENSOR 27, Bruker, Germany) and X-ray photoelectron spectroscopy (XPS; Escalab 250Xi, Thermo Fisher Scientific, USA) were utilized for further characterization. Fluorescence (FL) spectra were recorded using a fluorescence spectrometer (F-4700, Hitachi, Japan) equipped with a 150 W ozone-free xenon lamp, while ultraviolet–visible (UV–Vis) absorption spectra were obtained using a spectrophotometer (U-3900, Hitachi, Japan).

SDS-PAGE Gel electrophoresis

The protein content in Gd-CDs, HCM, and Gd-CDs@HCM was analyzed using sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) with a 12% pre-cast polyacrylamide gel. Total protein concentrations for each group were quantified using a bicinchoninic acid (BCA) protein assay kit, and protein solutions were diluted to a final concentration of 1 mg/mL. Electrophoresis was conducted in a Bio-Rad Mini-PROTEAN Tetra Cell system at 100 V for 1 h. After electrophoresis, the gels were stained with Coomassie Brilliant Blue and visualized using a gel documentation system.

Photothermal performance

A volume of 100 μL of Gd-CDs solutions at varying concentrations, along with ultrapure water as a control, was placed into 0.2 mL Eppendorf tubes. These samples were exposed to an 808 nm laser at a power density of 1 W/cm² for 5 min, during which temperature changes were recorded using an infrared thermal imaging camera. Subsequently, the temperature variation of a 0.2 mg/mL Gd-CDs solution under different laser power densities was similarly assessed. For the cyclic stability test, 100 μL of a 0.4 mg/mL Gd-CDs solution was irradiated with an 808 nm laser at 1 W/cm² for 5 min, followed by cooling to the initial temperature. This cycle was repeated five times, with temperature fluctuations documented via infrared thermal imaging. The photothermal conversion efficiency was calculated using the specified formula:

In this context, η represents the photothermal conversion efficiency, h denotes the thermal conductivity coefficient of the container, A signifies the surface area of the container, T_max refers to the maximum temperature of the solution, T_Sur represents the initial temperature of the solution, Q_s represents the heat loss, Q_s = (5.4 × 10^–4) I Js⁻¹, where I is the output power of the laser, and A_λ indicates the absorption value of the nanoplatform at 808 nm.

Additionally, a 0.4 mg/mL Gd-CDs solution, Gd-CDs@HCM solution, and ultrapure water were each placed into 0.2 mL centrifuge tubes. These samples were irradiated with an 808 nm laser at 1 W/cm² for 5 min, and temperature changes were recorded using an infrared thermal imaging camera.

Cell culture

HepG2 cells were cultured in MEM supplemented with 10% FBS. Hepa1-6 cells were cultured in DMEM supplemented with 10% FBS. THLE-2 cells were cultured in 1640 medium supplemented with 10% FBS. All cell lines were maintained at 37 ℃ in a humidified incubator with 5% CO₂.

Cytotoxicity assay

Healthy HepG2 and THLE-2 cells were enzymatically dissociated using trypsin and then resuspended in complete culture medium to a density of 1 × 10⁵ cells/mL. Each well of a 96-well plate was seeded with 100 μL of the cell suspension and incubated until adherence. Gd-CDs solutions at various concentrations (0, 12.5, 25, 50, 100, 200 μg/mL) were prepared, and cells were co-incubated with these solutions for 24 h. After incubation, cells were washed twice with PBS, and 100 μL of freshly prepared CCK-8 reagent was added to each well. Following a 1-h incubation, absorbance at 450 nm was measured using a multifunctional microplate reader, and the corresponding optical density (OD) values were recorded.

Hemolysis test

Fresh, healthy mouse blood was diluted with PBS and subjected to repeated centrifugation to prepare a red blood cell suspension. This suspension was incubated with varying concentrations of Gd-CDs@HCM solution samples for 4 h. Hemolytic activity was subsequently evaluated, with 1% Triton X-100 and PBS serving as the positive and negative controls, respectively.

Subcutaneous tumor-bearing mouse model

Male BALB/c and C57BL/6 J mice (4–6 weeks old) were obtained and acclimated for one week in a specific-pathogen-free (SPF) animal facility. HepG2 and Hepa1-6 cells in the logarithmic growth phase were harvested and suspended in MEM and DMEM culture media, respectively, to prepare cell suspensions at a concentration of 2 × 10⁶ cells/mL. A 0.2 mL aliquot of the cell suspension was subcutaneously injected into the right axillary region of BALB/c and C57BL/6 J mice to induce tumor formation. The experimental protocol was approved by the Animal Ethics Committee of Shanxi Province Cancer Hospital.

Bimodal imaging

FL imaging

HepG2 cells in good growth condition were resuspended in complete culture medium to achieve a concentration of 1 × 10⁴ cells/mL. A 2 mL aliquot was transferred to a specialized culture dish and incubated in a culture chamber for 12 h. The medium was then discarded, and the cells were washed with PBS. A medium containing 2% FBS was subsequently added, and the cells were starved for 1 h. Afterward, the medium was discarded, the cells were washed again with PBS, and complete culture media containing Gd-CDs or Gd-CDs@HCM (200 µg/mL) were added. The cells were incubated under the same conditions for 4 h, fixed with paraformaldehyde for 15 min, and imaged using confocal laser scanning microscopy (CLSM). For in vivo imaging, a Gd-CDs@HCM solution (10 mg/kg) was injected into HepG2 tumor-bearing mice, and FL signals were recorded at various time points using a small animal imaging system. BALB/c mice were euthanized at different post-injection intervals, and major organs (brain, heart, liver, spleen, lungs, kidneys, and bladder) were collected for ex vivo FL imaging to assess biodistribution.

MR imaging

HepG2 cells in good growth condition were cultured overnight in a 6-well plate. Gd-CDs@HCM solutions at different concentrations (0, 0.1, 0.2, 0.4, 0.8 mg/mL) were added to the wells and co-cultured for 4 h. After washing with PBS, the cells were digested with trypsin, centrifuged, and resuspended in 1% agarose solution for gelation. Imaging was performed using a small animal MR imaging system. For in vivo MR imaging, tumor-bearing mice were scanned with the same system. A Gd-CDs@HCM solution (10 mg/kg) was administered via tail vein injection, and scanning was conducted 6 h post-injection.

In vitro antitumor activity evaluation

The antiproliferative effects of Gd-CDs and Gd-CDs@HCM on HepG2 and THLE-2 cells were assessed using the CCK-8 assay. To further evaluate the targeted antitumor activity of Gd-CDs@HCM on HepG2 cells, additional in vitro experiments were conducted. HepG2 or THLE-2 cells were seeded into 96-well plates, with 5 replicate wells being set. Following overnight culture, cells were treated with controls, Gd-CDs, and Gd-CDs@HCM (200 µg/mL). After co-incubation for 4 h, cells in the Laser, Gd-CDs + Laser, and Gd-CDs@HCM + Laser groups were exposed to 808 nm laser irradiation at an intensity of 1 W/cm² for 5 min. The control group received no treatment. Subsequently, all groups were cultured for an additional 20 h. Absorbance values were determined by measuring the OD at 450 nm using a multifunctional microplate reader. To further assess the in vitro antitumor efficacy of Gd-CDs@HCM, cell viability and membrane integrity were evaluated using fluorescent dyes Calcein-AM and propidium iodide (PI) following the same experimental groups. Additionally, apoptosis rates in HepG2 cells were quantified using Annexin V/PI double staining and flow cytometry under the same treatment conditions.

Cell scratch assay

HepG2 cells in the logarithmic growth phase were seeded at a density of 2 × 10⁵ cells per well in a 6-well plate to ensure consistent seeding density and uniform cell distribution. After reaching 100% confluence, a sterile 200 μL pipette tip was used to create a vertical scratch, simulating a wound. Detached cells and debris were removed by washing the wells with PBS, and the old culture medium was replaced. Images of the scratched area were captured under a microscope immediately after scratching to serve as the 0-h control. The cells were then cultured in serum-free medium for 24 h. After incubation, the same scratched area was reexamined under the microscope, and images were taken to measure the wound width.

Cell invasion assay

The invasive capacity of HepG2 cells under different treatment conditions was evaluated using a cell invasion assay. Matrigel was evenly coated on the bottom surface of the Transwell upper chamber. HepG2 cells (2.5 × 10⁴ cells/well) were seeded in serum-free MEM medium in the upper chamber, while the lower chamber was filled with MEM medium supplemented with 10% FBS. The cells were incubated at 37 ℃ in a humidified atmosphere (5% CO₂ and 95% relative humidity) for 24 h. Following treatment under the specified experimental groups for 24 h, the cells were fixed with paraformaldehyde and stained with crystal violet solution. The number of invasive HepG2 cells was quantified using an inverted fluorescence microscope at 100 × magnification by counting cells in randomly selected fields.

Immunofluorescence

HepG2 cells were dissociated in complete culture medium to prepare a cell suspension at a concentration of 1 × 10⁴ cells/mL. A 2 mL aliquot of the suspension was transferred into a confocal culture dish using a pipette. Experimental groups were treated with Gd-CDs or Gd-CDs@HCM (200 µg/mL), while the control group received an equivalent volume of complete culture medium. After 4 h of incubation, cells in the experimental groups were exposed to laser irradiation at an intensity of 1 W/cm². Co-culture was maintained for 24 h. Subsequently, cells from all groups were collected, fixed with 4% paraformaldehyde, and blocked with 10% bovine serum albumin (BSA) for 1 h. The samples were incubated with primary antibodies for 2 h at room temperature, followed by incubation with fluorescently conjugated secondary antibodies for 1 h at room temperature. Nuclear staining was performed with DAPI for 10 min. Finally, the samples were visualized using confocal laser scanning microscopy (CLSM).

Detection of cellular CRT exposure via flow cytometry

HepG2 cells were seeded at a density of 3 × 10⁵ cells per well in a 6-well plate. Experimental groups were treated with appropriate concentrations of Gd-CDs or Gd-CDs@HCM, followed by a 4-h incubation. Cells were subsequently exposed to laser irradiation at an intensity of 1 W/cm², while the control group received an equivalent volume of complete culture medium. Co-culture was maintained for 24 h. Cells from each group were then collected, fixed with 4% paraformaldehyde, and incubated with primary antibodies for 1 h at room temperature. This was followed by incubation with fluorochrome-conjugated secondary antibodies for 30 min at room temperature. Finally, the samples were analyzed using flow cytometry.

Release of HMGB1 and ATP from cells

HepG2 cells were seeded at a density of 3 × 10⁵ cells per well in a 6-well plate. Experimental groups were treated with appropriate concentrations of Gd-CDs or Gd-CDs@HCM, followed by a 4-h incubation. Cells were then exposed to laser irradiation at an intensity of 1 W/cm², while the control group received an equivalent volume of complete culture medium. Co-culture was maintained for 24 h. The supernatants from the cell cultures were collected and analyzed using HMGB1 and ATP assay kits. The concentrations of HMGB1 and ATP in each group were quantified using a microplate reader.

In vitro maturation of dendritic cells (DCs)

Hepa1-6 cells were seeded into the upper chamber of a Transwell system (pore size: 0.4 μm) and co-cultured with Gd-CDs and Gd-CDs@HCM (200 µg/mL) for 4 h. Subsequently, the upper chamber was washed three times with PBS to remove unbound nanoparticles. The experimental group was subjected to irradiation with an 808 nm near-infrared laser (1 W/cm², 5 min), while the control group received no irradiation. Immature DCs, derived from mouse bone marrow cells, were seeded in the lower chamber of the Transwell system, with the treated Hepa1-6 cells placed in the upper chamber. Co-culture was maintained for 24 h at 37 ℃ and 5% CO₂ in RPMI-1640 complete culture medium supplemented with 10% FBS and 1% penicillin/streptomycin. Following co-culture, the DCs in the lower chamber were collected and incubated in the dark with anti-CD11c-FITC, CD80-PE, and CD86-APC antibodies for 30 min. Flow cytometry was employed to analyze the proportion of CD80⁺CD86⁺CD11c⁺ cells, thereby assessing the maturation status of the DCs.

Evaluation of antitumor efficacy in vivo

Tumor-bearing mice were administered physiological saline, Gd-CDs, or Gd-CDs@HCM via tail vein injection. After 12 h, photothermal imaging was performed using an 808 nm laser (1 W/cm²). For in vivo tumor treatment, subcutaneous tumor-bearing mice (tumor volume: 60–100 mm³) were randomly divided into six groups (n = 5 per group): (1) Saline, (2) Gd-CDs, (3) Gd-CDs@HCM, (4) Laser, (5) Gd-CDs + Laser, and (6) Gd-CDs@HCM + Laser. The injection volume for physiological saline was 100 µL, and the Laser group received 5 min of 808 nm laser irradiation (1 W/cm²). In the Gd-CDs and Gd-CDs@HCM groups, a dosage of 5 mg/kg of Gd-CDs was administered. Tumor dimensions (length and width) were measured every other day using calipers, and tumor volume was calculated using the formula:

where (a) is tumor length and (b) is tumor width. Body weight was also recorded. After 14 d of treatment, tumors were excised, photographed, and weighed. Tumor Growth Inhibition (TGI) was calculated using the formula:

RTV denotes relative tumor volume, V₀ is the tumor volume at day 0, and V_t is the tumor volume at day 14.

Cytokine and blood analysis

After 14 d of treatment, blood samples were collected from each group, and serum was extracted. Cytokine levels (IL-10, TNF-α, IL-6, and IFN-γ) were measured using ELISA kits. Additionally, complete blood count and biochemical analyses were performed.

Histological analysis

After 14 d of treatment, hearts, livers, spleens, lungs, kidneys, and tumor tissues were collected from each group. The tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and subjected to hematoxylin and eosin (H&E) staining. An experienced pathologist analyzed the stained tissue sections.

In vivo anti-tumor immune activation

After post-treatment for 14 d, tumor tissues, spleens, and inguinal lymph nodes were collected from each group. Immunofluorescence staining was performed on tumor tissues to evaluate the anti-tumor immune response in vivo. Additionally, tumor tissues, spleens, and inguinal lymph nodes were mechanically dissociated into single-cell suspensions by grinding the tissues until a turbid solution was obtained. The suspensions were filtered to remove connective tissue debris and incubated with 5% BSA for 15 min to block nonspecific binding. Cells were then stained with flow cytometry antibodies targeting the following populations:

- Mature DCs: CD11c-FITC, CD80-PE, CD86-APC.

- CD8⁺ T cells: CD45-PerCP, CD3- APC, CD8-PE.

-Tregs: CD45-PerCP, CD4-FITC, CD25-APC, Foxp3-PE.

The stained cells were incubated in the dark for 30 min and subsequently analyzed using FCM.

Statistical analysis

Statistical analyses were conducted using SPSS version 23.0, and results are expressed as mean ± standard deviation (SD). Statistical significance was assessed using one-way analysis of variance (ANOVA), with significance levels set at *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Author contributions

W.D. conceived and designed the study. W.D., Y.W., C.C., H.M. and S.Z. performed the experiments. W.D., Y.Y., and S.Y. analyzed the data. W.D. wrote the original draft. Y.X., L.C., Y.Y., and S.Y. reviewed and edited the manuscript. All authors have read and approved the final version of the manuscript.

Funding

This work was supported by the National Natural Science Foundation of China (82172048, U21A20378), the Shanxi Scholarship Council of China (2024–058, 2022–039), the Four “Batches” Innovation Project of Invigorating Medical through Science and Technology of Shanxi Province (2023XM012), the Science and Education Cultivation Fund of the National Cancer and Regional Medical Center of Shanxi Provincial Cancer Hospital (TD2023003, BD2023004, QH2023013), the Science and Technology Cooperation and Exchange Special Project of Shanxi Province (202304041101030, 202304041101002), the Shanxi Center of Technology Innovation for Controlled and Sustained Release of Nano-drugs (202104010911026), the Graduate Practice Innovation Project in Shanxi Province (2025SJ023).

Data availability

All data are available in the main text, supporting information, and are also on request from the corresponding author.

request from the corresponding author.

Declarations

Ethics approval and consent to participate

All animal experiments received approval from the Animal Ethics Committee of Shanxi Cancer Hospital (Approval Number: DWLL2025020).

Consent for publication

All authors have read and approved this version of the article for publication.

Competing interests

The authors declare no competing interests.