A layered target-release nanoparticle system for normalizing the lung cancer microenvironment and enhancing antitumor effect

Introduction

The biological characteristics of tumor tissue are markedly different from those of normal tissue. This is primarily reflected in abnormal angiogenesis, impaired blood flow, and increased permeability (1). Tumor blood vessels are the foundation for tumor growth and metastasis, and they are also crucial for the development and spread of tumors (2). Under normal circumstances, solid tumors that do not exceed 2 mm³ in volume mainly transport oxygen and nutrients to the deep layers of the tumor through simple diffusion (3). However, once the tumor reaches 2 mm³ in volume, cells become hypoxic, triggering rapid and disorderly proliferation of tumor blood vessels, resulting in a large number of blood vessels with large endothelial gaps and incomplete structures in the solid tumor tissue (3). The malformation of tumor blood vessels and the high permeability of the vessel walls significantly hinder the transport of nanoparticles (NPs) within the vessels (4). Moreover, tumor tissue lacks a functional lymphatic system, a deficiency that leads to excessive pressure differences from the tumor center to the periphery, hampering the transport of NPs through the vessels to the deep layers of the tumor (5). Furthermore, tumor tissue has a denser collagen fiber network and fibroblasts than does normal tissue, which can lead to increased interstitial pressure in the tumor, thereby compressing blood vessels, affecting blood flow, and impeding the transport of NPs (6). In summary, promoting the normalization of the tumor microenvironment is critical to improving the delivery of NPs in tumor tissue.

Ligustrazine (LT), a lipophilic active monomer component extracted from the traditional Chinese medicinal herb Chuanxiong, is one of the most effective components for improving the microcirculation within blood vessels (7,8). Recent research has shown that LT can significantly increase the integrity of tumor blood vessels, reduce tumor microvascular density, and promote the growth of healthy blood vessels, thereby normalizing tumor vasculature (9). The Chinese herb Leigongteng (Tripterygium wilfordii) is highly toxic and associated with the liver and kidney meridians. It possesses the effects of dispelling wind and dampness, reducing swelling and pain, and detoxifying and killing parasites (10,11). Numerous studies have demonstrated that triptolide (TPL), derived from the root bark of Leigongteng, exerts significant antiproliferative effects in various types of cancer cells, such as breast cancer cells (MCF-7 and MDA-MB-231), pancreatic cancer cells (MiaPaCa-2), and liver cancer cells (HepG2), making it a promising natural antitumor drug (12,13). The toxicity issue of TPL has also attracted considerable research attention. Encapsulating the drug in NPs to avoid direct exposure to the physiological environment is a common strategy for realizing detoxification (14,15).

The aim of this study was to enhance the antitumor efficacy of NPs by using TPL as an antitumor drug and LT as a microenvironment modulator for the synergistic treatment of patients with lung cancer. The newly developed composite lipid system consists of two-tiered NPs. The inner layer consists of chitosan and folic acid-modified chitosan (FA-CS), which act as carriers encapsulating TPL for the precise targeting of tumor cells. The outer layer contains pH-sensitive dioleoylphosphatidylethanolamine (DOPE) as the lipid material, encapsulating both LT and triptolide-loaded nanoparticles (TPL-NPs) to form LT@TPL-NPs. These composite NPs reach the tumor tissue through the enhanced permeability and retention (EPR) effect and release LT and TPL-NPs under low pH conditions. LT is responsible for normalizing the tumor microenvironment and synergistically facilitating the deep delivery of TPL-NPs into the tumor. TPL-NPs, modified with surface folic acid ligands and positively charged characteristics, are specifically taken up by deep tumor cells and ultimately release the drug into the cytoplasm. We present this article in accordance with the ARRIVE reporting checklist (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2026-1195/rc).

Methods

Preparation of LT@TPL-NPs

Chitosan (weight-averaged molecular weight, Mw ~70 kDa; degree of deacetylation >92%) was used as the base carrier material and reacted with folic acid-N-hydroxysuccinimide at a ratio of 10:1 to 1:1 (w/w) in dimethyl sulfoxide under anaerobic conditions for 48 h in the dark. After excess folic acid was removed via dialysis against running water, freeze-drying was applied to obtain FA-CS. With FA-CS serving as the primary material and sodium tripolyphosphate as an ionic cross-linking agent, TPL-NPs were prepared with a classic method that involved dissolution, coincubation, crosslinking, and lyophilization. DOPE was used as the lipid material. The film dispersion method was applied to dissolve the DOPE in chloroform to form a film. The film was hydrated under light-protected conditions with an aqueous solution of LT and TPL-NPs to synthesize LT@TPL-NPs, and the particle size was adjusted via ultrasonication.

Characterization of LT@TPL-NPs

To examine the pH-dependent disassembly of LT@TPL-NPs, the NPs were placed in phosphate-buffered saline (PBS) solution at pH values of 7.4, 6.8, and 5.0 for 2 h. The morphology of the NPs was captured via transmission electron microscopy (TEM). The particle size and zeta potential were analyzed via dynamic light scattering (DLS).

pH-response release of LT

LT@TPL-NPs were placed in PBS solutions with pH levels of 7.4, 6.8, and 5.0 at 37 ℃ for dialysis. At various time points, 2 mL of the solution was withdrawn and replenished with 2 mL of fresh PBS. The LT and TPL levels in the solution were measured via high-performance liquid chromatography (HPLC).

Internalization of LT@TPL-NPs by cells

Lung cancer cell lines A549 and H1299 were purchased from the Cell Bank, Chinese Academy of Sciences (Shanghai, China), and maintained in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) (Gibco, Thermo Fisher Scientific, Waltham, MA, USA). The internalization of NPs by these cells was examined with a confocal laser scanning microscope (CLSM; Leica, Wetzlar, Germany). For these studies, cells were plated at a density of 10⁵ per well in confocal dishes or six-well plates and cultured in DMEM with pH levels adjusted to 7.4 and 6.8; LT@TPL-NPs were added at a concentration of 20 µg/mL for durations of 4 and 12 h. Subsequently, the cells underwent fixation, permeabilization, and staining with 4′,6-diamidino-2-phenylindole (DAPI) prior to being analyzed with a CLSM.

A549 multicellular spheroid culture and internalization of NPs

A 1.5% (w/v) agarose solution prepared in DMEM was heated until the agarose was completely dissolved. Then, 80 µL of this solution was added into each well of a 96-well plate and allowed to cool and solidify, forming an agarose bed. Subsequently, cells were seeded into the 96-well plates at a density of 2×10³ per well and cultured in DMEM for a period of 6 days to allow them to develop into multicellular spheroids. These spheroids were then carefully transferred to confocal dishes and incubated in DMEM adjusted to a pH of 7.4 and 6.8. After exposure to LT@TPL-NPs at a dosage of 20 µg/mL for 4 and 12 h, the spheroids were rinsed, resuspended in fresh DMEM, and visualized with a CLSM.

Cell viability

To assess the micelles’ cytotoxic effects, cancer cells were plated in a 96-well plate at a density of 5×10³ cells per well and then exposed to the NPs for durations of 24, 48, and 72 h. Subsequently, the cells were incubated with a mixture of 200 µL of fresh medium and 20 µL of Cell Counting Kit 8 (CCK-8) reagent at 37 ℃ for an additional 1.5 h. The absorbance at 450 nm was then measured with a model 680 spectrophotometric plate reader (Bio-Rad Laboratories, Hercules, CA, USA) to determine the viability of the cells. In the EdU experiment, cells were prepared by fixation and permeabilization, which was followed by incubation with EdU staining solution (Beyotime Biotechnology, Shanghai, China) and DAPI (Sigma-Aldrich, St. Louis, MO, USA). Images were acquired with a fluorescence microscope (Leica).

Cell apoptosis

Cancer cells were grown in six-well plates to reach a confluence of 60–70% before being subjected to the NP treatment for a period of 24 h. Subsequent to treatment, the cells were collected through centrifugation and processed for staining with the Annexin V-fluorescein isothiocyanate/propidium iodide (Annexin V-FITC/PI) apoptosis detection kit (NeoBioscience, Beijing, China) as per the protocol provided by the manufacturer.

Cell migration and invasion

Cells were seeded in serum-free medium into the upper chamber of a Transwell insert (8-µm pore size membrane). The lower chamber was filled with medium containing 10% FBS. The plate was then incubated for 48 h to allow cells to migrate or invade through the membrane. After incubation, nonmigrated or noninvaded cells on the upper surface of the membrane were removed, and the cells that had traversed to the lower surface were fixed, stained with crystal violet (Beyotime), and counted under a microscope to quantify migration and invasion.

Tube formation assay

Human umbilical vein endothelial cells (HUVECs) were seeded in a 96-well plate coated with Matrigel (Corning Inc., Corning, NY, USA) with approximately 2.5×10⁴ cells per well and incubated at 37 ℃ in a CO₂ incubator for 4 to 16 h to allow the cells to form tube-like structures. After incubation, the images of tubes were captured under a microscope and quantified via ImageJ software (ImageJ, US National Institutes of Health, Bethesda, MD, USA).

Western blot assay

The cells were washed with PBS and lysed in RIPA buffer containing protease inhibitors. Subsequently, 20 µg of protein was separated on 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel and transferred to nitrocellulose membranes in transfer buffer (Boston BioProducts, Ashland, MA, USA) containing 20% methanol. Apoptosis-related proteins were detected via treatment with rabbit monoclonal anti-Bcl-2, anti-Bax, and anti-caspase-3, along with a subsequent treatment with secondary horseradish peroxidase (HRP)-conjugated antibodies. Detection was conducted on a chemiluminescence imaging system (Tanon 5200, Tanon Science & Technology Co., Ltd., Shanghai, China).

In vivo antitumor assay

All animal experiments were performed under a project license (Approval No. DWSY-2024-007) granted by the Animal Care and Use Committee of The Third Affiliated Hospital of Wenzhou Medical University, in compliance with the institutional guidelines for the care and use of laboratory animals. Female severe combined immunodeficiency (SCID) nude mice (6–8 weeks old, body weight 18–22 g) were purchased from Shanghai SLAC Laboratory Animal Co., Ltd. (Shanghai, China) and housed under specific pathogen-free conditions with a 12 h light/dark cycle, with free access to food and water.

A549 cells (1×10⁶ cells in 100 µL of PBS) were subcutaneously injected into the right flank of each mouse. When the tumor volume reached approximately 100 mm³ (designated as day 0), the mice were randomly assigned to three treatment groups (n=6 mice per group): (I) PBS (control), (II) TPL-NPs, and (III) LT@TPL-NPs.

All treatments were administered via tail vein injection at a dose of 20 mg TPL-equivalent per kg body weight in a volume of 100 µL per mouse. The injections were given twice per week (Monday and Thursday) for two consecutive weeks (total of four doses). Tumor dimensions were measured every 3 days using a digital caliper, and tumor volume was calculated using the formula: volume (mm³) = (length × width²) × 1/2.

On day 14 after the first treatment, the mice were euthanized by cervical dislocation under deep anesthesia (isoflurane overdose). Tumors were excised, weighed, photographed, and immediately fixed in 4% paraformaldehyde for subsequent histological examination.

Immunohistochemical (IHC) staining

IHC analysis was carried out as per established methods. Tumor tissues were routinely deparaffinized, underwent citrate-based antigen retrieval, and were treated with hydrogen peroxide to quench endogenous peroxidase activity. The sections were then incubated with 10% bovine serum albumin (BSA; Beyotime) at room temperature for 1 h to block nonspecific binding. Subsequently, the sections were exposed to a primary antibody specific for CD31 at 4 ℃ overnight. On the subsequent day, they were treated with an HRP-conjugated secondary antibody (rabbit) for 45 minutes at room temperature. Immunoreactive sites were detected with diaminobenzidine substrate (Beyotime), and the sections were counterstained with hematoxylin. The slides were examined and imaged with a microscope (Leica). For quantitative evaluation of vascular normalization, five non-overlapping fields of view were randomly captured per section under unified magnification. ImageJ software was used to calculate the CD31-positive area fraction, which was defined as microvessel density (MVD).

Masson trichrome staining

To measure the collagen composition and extracellular matrix (ECM) remodeling, sections of tumor tissues were processed with a Masson trichrome staining kit (Solarbio, Beijing, China) in accordance with the manufacturer’s guidelines. The staining kit’s protocol yielded distinct colorations: fibrous tissues such as collagen and reticular fibers appeared blue; nuclei appeared purple-black; and cytoplasm, muscular tissue, and red blood cells appeared red. For quantitative analysis of collagen deposition, five random visual fields were photographed for each slice, and the percentage of blue collagen fiber area relative to total tissue area was calculated via ImageJ software to assess ECM remodeling.

Biosafety analysis

Prior to being embedded in paraffin, major organ tissues (heart, liver, spleen, lung, kidney) underwent dehydration through a graded ethanol series, which was followed by treatment with xylene. Tissue sections, each 10-µm thick, were prepared. Postdeparaffinization, these sections were stained with hematoxylin and eosin (HE) staining solution for histological evaluation of tissue necrosis, inflammatory infiltration and organ injury. Microscopic examination was then conducted to capture images of the stained slides. For systemic toxicity assessment, peripheral blood was collected from all experimental mice for serum biochemical analysis. The levels of total protein (T-Pro), albumin (ALB), total bilirubin, alanine aminotransferase (ALT), creatinine (CRE), blood urea nitrogen (BUN), and uric acid (UA) were measured with a biochemical analyzer, focusing on liver and kidney function, the primary organs damaged by free TPL systemic exposure.

Statistical analysis

Data are expressed as the mean ± standard deviation. Statistical evaluations were conducted with GraphPad Prism version 5 software (Dotmatics, Boston, MA, USA). For pairwise comparisons, a Student’s t-test was employed, while one-way analysis of variance coupled with post hoc multiple comparisons was applied to assess differences across multiple groups. A P value <0.05 indicated statistical significance.

Results

Characterization and pH-responsive disassembly of LT@TPL-NPs

To investigate the pH-dependent transition of LT@TPL-NPs, the NPs were incubated in PBS solutions at pH 7.4, 6.8, and 5.0 for 2 h. TEM observations revealed that LT@TPL-NPs exhibited a uniform spherical morphology at pH 7.4. At pH 6.8, the particle size was significantly reduced and the bilayer structure became loose. At pH 5.0, the spherical structure was almost completely disintegrated (Figure 1A). DLS analysis confirmed the pH-dependent alterations in particle size distribution (Figure 1B). The zeta potential increased from approximately 10 mV at pH 7.4 to about 20 mV at pH 6.8, and further increased to nearly 30 mV at pH 5.0 (Figure 1C).

Figure 1 In vivo antitumor efficacy and biosafety of LT@TPL-NPs. Therapeutic effects and microenvironment remodeling in A549 xenograft models. (A) Representative images of excised tumors from different groups. (B) Tumor volume growth curve during treatment. Mean ± SD. *P<0.05, **P<0.01 vs. TPL-NPs group. (C) Tumor weight at the end of treatment. Mean ± SD. *P<0.05; **P<0.01. (D) Immunohistochemical staining of CD31 for quantitative microvessel density analysis, a representative marker of tumor vascular normalization. Scale bar = 100 μm. (E) Masson trichrome staining for quantitative detection of collagen fiber area fraction to evaluate extracellular matrix remodeling. Collagen fibers: blue; nuclei: black-brown. Scale bar = 100 μm. (F) Hematoxylin and eosin (HE) staining of major organs (heart, liver, spleen, lung, kidney). Scale bar = 100 μm. TPL, triptolide; LT, ligustrazine; PBS, phosphate-buffered saline; IHC, immunohistochemistry; HE, hematoxylin and eosin.

To evaluate the pH-responsive release behavior, the cumulative release of LT and TPL from LT@TPL-NPs was measured via HPLC. As shown in Figure 1D, both LT and TPL exhibited minimal release at pH 7.4, while rapid and significant release was observed under acidic conditions (pH 6.8 and 5.0), confirming pH-triggered drug release.

Enhanced penetration and cellular uptake of LT@TPL-NPs at acidic pH

In A549 multicellular tumor spheroids, LT-FITC@TPL-Cy5.5-NPs were predominantly confined to the peripheral layer at pH 7.4, with negligible fluorescent signals observed in the spheroid core after 12 h of incubation. By contrast, under acidic pH 6.8 conditions, both green LT-FITC and red TPL-Cy5.5 fluorescence signals penetrated the whole spheroid as early as 4 h, and signal intensity was further elevated at 12 h (Figure 3A).

Figure 3 In vitro antitumor effects of LT@TPL-NPs. Inhibition of proliferation, apoptosis, migration, invasion, and angiogenesis by LT@TPL-NPs. (A) Cell viability of A549 and H1299 cells after treatment with PBS, TPL-NPs, or LT@TPL-NPs for 24, 48, and 72 h (CCK-8 assay). Mean ± SD, *P < 0.05, **P < 0.01 vs. TPL-NPs group. (B) Colony formation of A549 and H1299 cells. Additional functional assays including EdU staining, flow cytometry apoptosis detection, Transwell invasion and western blot analysis are displayed in Figure 2C-2F. Footnote: TPL, triptolide; LT, ligustrazine; PBS, phosphate-buffered saline; CCK-8, Cell Counting Kit-8; EdU, 5-ethynyl-2′-deoxyuridine; DAPI, 4′,6-diamidino-2-phenylindole; FITC, fluorescein isothiocyanate; PI, propidium iodide; HUVECs, human umbilical vein endothelial cells.

CLSM imaging of A549 and H1299 cells confirmed efficient intracellular internalization of LT@TPL-NPs at both physiological pH 7.4 and tumor-mimicking acidic pH 6.8. Notably, cellular uptake efficiency was markedly augmented within the acidic microenvironment relative to neutral culture conditions (Figure 3B). No additional phenotypic or molecular functional panels C-F are included in Figure 3, so all related quantitative functional data are presented separately in Figure 2.

Figure 2 Tumor penetration and cellular internalization of LT@TPL-NPs. Enhanced penetration and uptake of LT@TPL-NPs in acidic tumor-mimicking microenvironment. (A) Fluorescence distribution of LT-FITC@TPL-Cy5.5-NPs in A549 multicellular tumor spheroids at pH 7.4 and 6.8 for 4 and 12 h. Green: LT-FITC; red: TPL-Cy5.5. Scale bar = 100 μm. (B) Confocal laser scanning microscopy images of cellular internalization in A549 and H1299 cells at pH 7.4 and 6.8 for 4 h. Blue: DAPI nuclear staining; green/red: nanoparticle signals. Scale bar = 50 μm. LT, ligustrazine; FITC, fluorescein isothiocyanate; TPL, triptolide; Cy5.5, a near-infrared fluorescent dye.

LT@TPL-NPs inhibit proliferation, migration, invasion and angiogenesis in lung cancer cells

CCK-8 assays showed that LT@TPL-NPs significantly reduced the viability of A549 and H1299 cells at 24, 48, and 72 h in a time-dependent manner, with stronger inhibitory effects than TPL-NPs alone (Figure 2A). Colony formation assays revealed that LT@TPL-NPs markedly decreased the number and size of colonies formed by lung cancer cells (Figure 2B). EdU assays further confirmed that LT@TPL-NPs significantly suppressed the proliferation of A549 and H1299 cells (Figure 2C).

Flow cytometry demonstrated that LT@TPL-NPs induced a markedly higher apoptosis rate in lung cancer cells compared with PBS and TPL-NPs groups (Figure 2D). Western blot analysis showed that LT@TPL-NPs significantly downregulated the expression of anti-apoptotic Bcl-2 and upregulated the levels of pro-apoptotic Bax and cleaved caspase-3 (Figure 2E).

Transwell assays indicated that LT@TPL-NPs significantly reduced the number of migrated and invaded A549 and H1299 cells compared with other groups. Tube formation assays showed that LT@TPL-NPs obviously decreased the number of complete tube-like structures formed by HUVECs, indicating suppressed angiogenesis (Figure 2F).

LT@TPL-NPs suppress tumor growth and remodel tumor microenvironment in vivo

In A549 xenograft nude mice, TPL-NPs moderately reduced tumor volume and weight. In contrast, LT@TPL-NPs exerted significantly stronger inhibitory effects on tumor growth, with much smaller tumor volume and lower tumor weight compared with PBS and TPL-NPs groups (Figure 4A-4C).

Figure 4 Characterization and pH-responsive disassembly of LT@TPL-NPs. pH-triggered structural changes and drug release behavior of LT@TPL-NPs. (A) Transmission electron microscopy images of LT@TPL-NPs at pH 7.4, 6.8, and 5.0 for 2 h. Scale bar = 200 nm. (B) Dynamic light scattering analysis of particle size distribution under different pH conditions. (C) Zeta potential changes of LT@TPL-NPs at pH 7.4, 6.8, and 5.0. (D) Cumulative release of LT and TPL from LT@TPL-NPs at different pH values, measured by HPLC, indicating pH-dependent drug release. HPLC, high-performance liquid chromatography; LT, ligustrazine; LT@TPL-NPs, ligustrazine and triptolide co-loaded nanoparticles; TPL, triptolide.

We performed quantitative analysis on two core indicators of tumor microenvironment normalization: CD31 MVD for vascular normalization and Masson collagen area fraction for ECM remodeling. IHC staining and quantitative statistics showed that LT@TPL-NPs markedly reduced CD31-positive area fraction, confirming effective vascular normalization by pruning disordered pathological microvessels. Masson trichrome staining and quantitative calculation demonstrated that LT@TPL-NPs significantly lowered the proportion of collagen fibers in tumor stroma, alleviating excessive ECM deposition and interstitial pressure to achieve collagen matrix remodeling. HE staining of major organs including heart, liver, spleen, lung, and kidney showed no obvious histological damage, tissue necrosis or inflammatory infiltration in the LT@TPL-NPs group. All tested serum liver and kidney function biochemical indices (ALT, total bilirubin, CRE, BUN, UA, ALB, T-Pro) of mice treated with LT@TPL-NPs remained within normal physiological ranges. Consistent with previous literature reports, free TPL induces significant elevation of liver and kidney injury markers and obvious pathological lesions in visceral organs due to non-specific systemic distribution. These preliminary safety data confirm that the bilayer NP delivery system effectively alleviates the systemic toxicity of TPL.

Discussion

Abnormal tumor vasculature and dense ECM are major barriers that restrict drug penetration and impair therapeutic efficacy in solid tumors (1,4,5). In this study, we constructed a pH-responsive bilayer NP system (LT@TPL-NPs) for sequential delivery of LT and TPL, aiming to normalize the tumor microenvironment and enhance antitumor efficiency.

The LT@TPL-NPs platform was designed to achieve spatiotemporally controlled drug release in response to tumor acidity. At pH 6.8 (tumor microenvironment), the outer pH-sensitive lipid layer composed of DOPE disassembled and released LT first, while the inner folate-modified chitosan core was exposed with reduced size and increased positive potential, which facilitated deep tumor penetration and cellular uptake (16). In the endosomal acidic environment (pH ≈ 5.0), the inner core further decomposed to release TPL. This hierarchical disassembly and sequential release pattern is conducive to the synergistic action of microenvironment regulation and targeted chemotherapy, which is consistent with the design concept of intelligent tumor-targeted delivery systems (14).

LT, a natural monomer derived from Chuanxiong, has been verified to improve microcirculation and normalize tumor blood vessels (7,9). Our results confirmed that LT released from LT@TPL-NPs reduced CD31-positive microvessels and collagen deposition, thereby remodeling the ECM and alleviating high interstitial pressure. These changes improved the permeability and penetration depth of inner TPL-NPs into deep tumor tissue, which is critical for overcoming drug delivery barriers in solid tumors.

Quantitative detection of two core indicators further confirmed LT-mediated dual remodeling of tumor vasculature and collagen matrix. CD31-based MVD serves as a gold-standard indicator to judge vascular normalization; the reduced CD31-positive area in LT@TPL-NPs group indicated that LT eliminated redundant malformed angiogenic vessels and restored relatively functional, normalized tumor vasculature. Meanwhile, the decreased collagen fiber area fraction from Masson staining reflected relieved dense stromal barriers and lowered intratumoral interstitial fluid pressure. Combined with in vitro HUVEC tube formation results that directly reflect endothelial angiogenic capacity, these in vivo quantitative data jointly verified that LT simultaneously realizes vascular normalization and collagen remodeling, fundamentally removing the physical barriers restricting deep NP penetration into solid tumors.

In addition, although positively charged TPL-NPs achieve efficient intracellular uptake within acidic tumor tissue, bare cationic NPs encounter severe systemic delivery defects in vivo. Unshielded cationic nanocarriers readily adsorb abundant plasma opsonin proteins to form a protein corona, which triggers rapid clearance by reticuloendothelial system (RES) macrophages in the liver and spleen, shortening blood circulation time and reducing tumor accumulation. Moreover, positive NPs electrostatically bind negatively charged erythrocyte membranes, disrupting membrane integrity and inducing hemolysis and systemic blood toxicity. Our bilayer pH-responsive LT@TPL-NPs overcome these drawbacks via a charge-shielding strategy. At physiological pH 7.4 in blood, the outer DOPE lipid layer fully encapsulates the cationic inner core, and intact LT@TPL-NPs only exhibit weak positive zeta potential (~+10 mV). This weak surface charge markedly inhibits nonspecific plasma protein adsorption, RES-mediated clearance and hemolytic risk during systemic circulation. Only after accumulating in acidic tumor microenvironment (pH 6.8), the outer pH-sensitive lipid shell disassembles to release LT, locally exposing highly cationic TPL-NPs to strengthen electrostatic binding with tumor cell membranes and boost cellular uptake. Consistent with this design, in vivo histological and serum biochemical safety assays confirmed no obvious organ injury or hematological abnormalities in LT@TPL-NPs-treated mice, demonstrating that the hierarchical charge-shield/tumor-activated charge strategy balances long-circulation biosafety and intratumoral delivery efficiency.

The sequential disassembly property of LT@TPL-NPs enables a distinctive two-stage cascade synergistic therapeutic mechanism between LT and TPL. Once accumulated within the acidic tumor microenvironment, the outer pH-sensitive DOPE lipid shell decomposes and releases LT in advance. As a tumor microenvironment regulator, LT reduces MVD and collagen deposition, relieves high interstitial fluid pressure, and dismantles dense ECM barriers. Such microenvironment remodeling removes the major physical obstacles for NP penetration, allowing the subsequently exposed TPL-NPs to infiltrate into deep tumor layers and achieve high intratumoral drug retention. After deep intratumoral enrichment, folate-functionalized positively charged TPL-NPs are efficiently internalized by folate receptor-overexpressing lung cancer cells, and released TPL further suppresses multiple malignant phenotypes and triggers tumor cell apoptosis via mitochondrial pathways. This “microenvironment preconditioning first, targeted chemotherapy second” cascade synergy cannot be achieved by simply mixing free LT and free TPL, because free drugs lack tumor-specific accumulation and ordered spatiotemporal release. The combined barrier-remodeling effect of LT and direct cytotoxicity of TPL together produce remarkably stronger antitumor activity than single-agent treatment, which accounts for the superior therapeutic performance of LT@TPL-NPs observed both in vitro and in vivo.

TPL is a natural product with potent broad-spectrum antitumor activity (12,13). Our in vitro experiments showed that TPL released from LT@TPL-NPs significantly inhibited proliferation, migration and invasion of lung cancer cells, and induced apoptosis via the mitochondrial pathway by downregulating Bcl-2 and upregulating Bax and cleaved caspase-3. Notably, TPL has been reported to induce GSDME-mediated pyroptosis in head and neck cancer (23), suggesting that pyroptosis may also participate in the antitumor mechanism of TPL in lung cancer, which deserves further investigation.

In vivo, LT@TPL-NPs exhibited stronger tumor inhibition than TPL-NPs alone, confirming the synergistic effect between LT-mediated microenvironment normalization and TPL-mediated cytotoxicity. Meanwhile, no obvious organ injury or blood biochemical abnormalities were observed, indicating that the NP encapsulation strategy effectively reduced the potential toxicity of free TPL (15) and improved the biosafety of the therapeutic system.

Free TPL is well documented to trigger severe systemic hepatotoxicity and nephrotoxicity, which greatly restricts its translational application in antitumor therapy. Our bilayer pH-responsive LT@TPL-NPs relieve TPL-related systemic toxicity through two major design strategies. First, LT@TPL-NPs accumulate selectively in tumor tissue via the EPR effect and release drugs only under the acidic intratumoral microenvironment, which remarkably reduces non-specific TPL distribution in normal visceral organs. Second, the integrated lipid-chitosan bilayer isolates TPL from direct contact with circulating normal tissues during systemic delivery. In line with this design, in vivo HE staining of key organs exhibited no pathological lesions, and all serum liver and kidney biochemical indicators stayed at normal levels after LT@TPL-NPs administration. These preliminary biosafety results demonstrate that our nanoplatform can effectively mitigate the systemic toxic risks of free TPL while retaining strong synergistic antitumor potency.

This study has some limitations. First, only PBS and TPL-NPs groups were included for in vivo antitumor assessment, without free LT monotherapy or free LT plus free TPL combination groups. Although we supplemented comprehensive in vitro comparison data of free LT, free TPL and free LT + TPL mixture to preliminarily verify the unique synergistic superiority of sequential pH-triggered delivery of LT@TPL-NPs, the absence of corresponding in vivo data may limit the full interpretation of in vivo synergistic therapeutic effects. We have clearly acknowledged the lack of in vivo groups treated with free single drugs or free drug combinations as a major limitation of the present study. We also propose that follow-up standalone animal studies incorporating a full panel of control groups will be conducted in our future work to further corroborate our findings. Second, in vivo biodistribution experiments were not performed to directly visualize the tumor enrichment and in vivo pH-responsive release characteristics of LT@TPL-NPs. We will conduct systematic in vivo fluorescence distribution detection in subsequent research to fill this data gap. Third, the antitumor efficacy was only evaluated in subcutaneous xenograft models; orthotopic or genetically engineered lung cancer models are needed to better simulate clinical pathophysiological features. Fourth, the impact of LT@TPL-NPs on the immune microenvironment, such as dendritic cell maturation and T cell infiltration, was not explored. Fifth, the long-term stability, pharmacokinetics and biodistribution of LT@TPL-NPs require further optimization.

Conclusions

In this study, we successfully constructed a pH-responsive bilayer NP system (LT@TPL-NPs) for sequential targeted delivery of LT and TPL in lung cancer. The NPs achieve hierarchical disassembly in the acidic tumor microenvironment, releasing LT first to normalize tumor blood vessels and remodel the ECM, thereby eliminating physical barriers and enhancing deep tumor penetration of TPL-NPs (7,9). The inner folate-modified core subsequently releases TPL to induce cancer cell apoptosis and suppress proliferation, migration, invasion, and angiogenesis (21,22,23). The two-stage cascade action of LT and TPL generates prominent synergistic antitumor efficacy both in vitro and in vivo with favorable biosafety, effectively overcoming the common drug delivery barriers in solid tumors (1,14,16).

This nanoplatform provides a promising and translatable strategy for enhanced therapy of lung cancer and other solid tumors with an acidic microenvironment.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE reporting checklist. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2026-1195/rc

Data Sharing Statement: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2026-1195/dss

Peer Review File: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2026-1195/prf

Funding: This study was supported by the Medical and Health Research Project of Zhejiang Province (No. 2024KY400) and the Beijing Science and Technology Innovation Medical Development Foundation (No. KC2023-JX-0186-PQ011).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2026-1195/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. Animal experiments were performed under a project license (Approval No. DWSY-2024-007) granted by the Animal Care and Use Committee of The Third Affiliated Hospital of Wenzhou Medical University, in compliance with the institutional guidelines for the care and use of laboratory animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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(English Language Editor: J. Gray)