A Ferrous‐Supply‐Regenerating Lipid Nanoparticle Integrating RNAi Induces Ferroptosis for Cancer Therapy

ABSTRACT

Targeting iron‐dependent ferroptosis with siRNA has emerged as a promising strategy for cancer treatment. The concept of ferrous‐supply regeneration, inspired by electro‐Fenton technology, has also gained interest as a non‐apoptotic approach to induce ferroptosis. However, integrating both strategies into a single lipid nanoparticle (LNP) remains challenging. To address this, we developed ALSF, a novel LNP in which DSPC is replaced with arachidonic acid (AA) to enable co‐delivery of Glutathione Peroxidase 4‐siRNA (GPX4‐siRNA) and Fe^{3 +}. ALSF facilitates acid‐ and H₂O₂‐dependent iron recycling, continuously regenerating Fe^{2 +} to drive ROS production, while GPX4‐siRNA silences GPX4 to induce lipid peroxidation (LPO) and ferroptosis. Elevated H₂O₂ levels in cancer cells promote selective iron recycling, lysosomal disruption, and siRNA escape, enabling effective gene knockdown at lower doses. In vivo, ALSF suppressed tumor growth in a C918 xenograft model with favorable biosafety. This study is the first to integrate a ferrous‐supply‐regeneration system with siRNA into a single LNP platform, offering a selective ferroptosis‐inducing strategy for ocular cancers and other ferroptosis‐sensitive malignancies.

Short abstract

Targeting iron‐dependent ferroptosis via siRNA and ferrous regeneration are promising anticancer strategies, yet their co‐encapsulation in LNPs is challenging. We constructed AA‐modified ALSF LNPs co‐delivering GPX4‐siRNA and Fe³⁺ to sustain Fe²⁺ supply and block GPX4. ALSF triggered selective ferroptosis and suppressed ocular tumor growth with good biosafety.

1. Introduction

Cancer poses a major clinical challenge, threatening patients' lives and, in certain cases, essential physiological functions [1]. The core dilemma in treatment lies in achieving tumor eradication without compromising vital functions—a goal that remains unattainable with current modalities (surgery, radiation, and chemotherapy) [1, 2, 3, 4]. Chemotherapy primarily relies on inducing caspase‐dependent apoptosis [5, 6]. However, frequent resistance to apoptosis in malignant cells severely limits efficacy and drives recurrence and metastasis [7, 8, 9]. Thus, novel cell death strategies that bypass apoptosis are urgently needed to address this clinical challenge [10, 11, 12], particularly for cancers that are insensitive to apoptotic pathways, including ocular cancers [13, 14].

Ferroptosis, an iron‐dependent cell death driven by LPO accumulation, offers a promising strategy for treating apoptosis‐resistant cancers like retinoblastoma by exploiting tumor metabolic vulnerabilities [15]. This process is governed by the “GPX4‐Fe‐LPO” axis: disruption of System Xc⁻ or GPX4 inactivates antioxidant defenses, while elevated iron catalyzes Fenton reaction‐driven LPO, ultimately destroying membrane integrity [16]. Given that Fe^{2 +} exhibits orders‐of‐magnitude higher catalytic activity than Fe^{3 +}, and intracellular Fe^{2 +} pools are tightly regulated, elevating Fe^{2 +} levels represents a key strategy for ferroptosis therapy [17, 18, 19, 20]. Inspired by electro‐Fenton technology, where cathodic Fe^{3 +} reduction enables sustained ROS generation via Fe^{2 +} regeneration, scientists propose introducing an analogous iron supply‐regeneration system into tumors to overcome insufficient Fe^{2 +} availability and potentiate ferroptosis [21].

Inspired by Yang et al., we noted that their nanoparticles co‐encapsulating Fe^{3 +} and unsaturated‐lipid‐rich phosphatidylcholine (PC) generate a Fe^{3 +}/Fe^{2 +} redox pair that directly catalyzes peroxidation of the lipid components, thus achieving Fenton‐reaction‐independent ferroptosis [22]. Beyond this class of unsaturated lipids, we have also focused on AA, a polyunsaturated fatty acid (PUFA) and the most representative ω‐6 PUFA, due to its unique role in ferroptosis. Structurally, AA contains four cis double bonds (C5–C6, C8–C9, C11–C12, C14–C15), rendering the bis‐allylic carbons highly vulnerable to oxidation. Oxidation of AA generates lipid radicals (L•), lipid peroxyl radicals (LOO•), and lipid hydroperoxides (LOOH), thereby propagating radical chain reactions [23, 24]. Importantly, this oxidative process also enables Fe³⁺ reduction to Fe²⁺ via radical‐mediated redox cycling, sustaining Fenton/Fenton‐like reactions and amplifying ROS production—a hallmark event in ferroptosis. Consequently, AA functions both as a major substrate for LPO and as a reducing agent capable of regenerating Fe²⁺ from Fe³⁺ [25, 26, 27].

Notably, free AA must first be esterified into membrane phospholipids—primarily phosphatidylethanolamine (PE)—via the ACSL4/LPCAT3 pathway to serve as a direct executor of LPO. Once incorporated, AA‐containing phospholipids, particularly AA‐PE, undergo iron‐dependent peroxidation to form AA‐derived phosphatidylethanolamine hydroperoxides (AA‐PE‐OOH) [28, 29], which have been identified as key death signalling molecules in ferroptosis. Enhanced AA metabolism and subsequent oxidation of AA‐containing phospholipids have been shown to markedly increase cellular susceptibility to ferroptosis. Based on these insights, we propose that co‐delivery of Fe^{3 +} and AA into tumour tissues—followed by lysosomal internalisation—could achieve a dual‐pronged effect: on one hand, it ensures an exogenous and sustained supply of Fe^{2 +} to sustain Fenton chemistry; on the other, it furnishes abundant LPO substrates in the form of AA‐containing membrane phospholipids, thereby efficiently potentiating ferroptosis.

We also noted an interesting phenomenon: compared with DSPC, the conventional structural lipid in standard LNPs for nucleic acid delivery [30], AA exhibits similar structural properties but superior functional attributes. Structurally, AA can replace DSPC as a structural component of LNPs owing to its hydrophobic chain length, which is comparable to that of DSPC tails, and its favorable affinity for lipid bilayers [31, 32, 33]. Functionally, AA provides iron‐coordinating capability that DSPC lacks, thereby offering the potential to transform LNPs from inert delivery carriers into therapeutic functional materials [34, 35, 36].

Therefore, in this study, we engineered a novel LNP (ALNP) by replacing the conventional structural lipid DSPC in standard LNPs with AA. The resulting formulation was co‐loaded with GPX4‐targeting siRNA (GPX4‐siRNA) and ferric ions (Fe^{3 +}) to generate ALNP@GPX4‐siRNA‐Fe^{3 +} (ALSF). Under neutral physiological conditions, Fe^{3 +} forms stable coordination complexes with carboxyl groups of AA or phosphate groups of nucleic acids, ensuring nanoparticle integrity [37, 38]. Upon internalization into cancer cell lysosomes, the acidic environment protonates these groups, triggering Fe^{3 +} dissociation and structural destabilization, which facilitates the rapid release of AA and siRNA [39, 40].

Tumor cells harbor elevated levels of H₂O₂ compared to normal cells, a feature exploited by our design [41]. The released AA engages in an iron cycling process with Fe^{3 +}, driving sustained Fe²⁺ regeneration and subsequent Fenton reaction‐mediated ROS production [42, 43]. These ROS not only initiate LPO but also promote lysosomal membrane disruption, thereby enhancing cytosolic delivery and the silencing efficacy of GPX4‐siRNA [44, 45, 46]. By concurrently targeting three pivotal nodes of the cellular redox system, namely GPX4, iron availability, and LPO, ALSF selectively induces ferroptosis in cancer cells while exhibiting minimal toxicity in normal cells with low H₂O₂ levels. This work presents a pathology‐dependent, tumor microenvironment‐responsive nanoplatform for safe and effective ferroptosis‐based cancer therapy.

2. Results and Discussion

2.1. Preparation and Characterization of ALSF

The formulation and preparation of LNP are well‐established at the laboratory scale, with defined component paradigms and standardized procedures [47]. In this study, we referenced the publicly available standard formulation of the approved drug Patisiran (Onpattro). SM‐102 was used as the ionizable lipid, and AA was employed as a structural lipid to replace DSPC [48]. The final formulation consisted of SM‐102:AA:cholesterol:PEG2000‐DMG at a molar ratio of 50:10:38.5:1.5. The N/P ratio for nucleic acid encapsulation was set at 6.

Both the 0:0 group (ALNP) and the 1:0 group (ALS) without Fe³⁺ showed poor size distribution after mixing, with polydispersity index (PDI) values above 0.2 (Figure 2A, B). When Fe³⁺ is introduced into the ethanol phase, it can form complexes with AA. During this reaction stage, the amino groups of SM‐102 may abstract protons from the carboxylic acid moieties of AA and trigger the ionization of AA. This proton transfer process facilitates the complexation between carboxylate anions dissociated from AA and Fe³⁺. Ultraviolet–visible (UV–vis) spectroscopic characterizations corroborate the above conclusion. The spectroscopic data reveal that SM‐102 exerts no inhibitory effect on the coordination between AA and Fe³⁺, and to a certain extent, promotes such coordination behavior (Figure S1). When the organic and aqueous phases are mixed via a microfluidic device to formulate ALSF, the abundant phosphate groups provided by siRNA become available, and Fe³⁺ may coordinate with either the carboxyl groups of AA or the phosphate groups of siRNA, thereby enhancing the stability of the resulting nanoparticles (Figure 1) [49, 50, 51]. ALSF formulations were prepared with varying GPX4‐siRNA:Fe³⁺ mass ratios (Figure 2A, B). Within the tested range (0:0 to 1:1, w/w), Fe³⁺ addition decreased PDI and increased average particle size. Specifically, the 1:0.1 and 1:0.33 groups showed PDI values of 0.137 ± 0.019 and 0.155 ± 0.026, respectively. The 1:0.66 and 1:1 groups further decreased PDI to below 0.1 (0.042 ± 0.014 and 0.074 ± 0.054), with the 1:0.66 group showing optimal uniformity. Additionally, Fe³⁺ may replenish intracellular iron in cancer cells, potentially promoting ferroptosis [52, 53, 54, 55]. However, after ultrafiltration and buffer exchange, the 1:1 group aggregated due to excessive Fe^{3 +}, resulting in a larger particle size and PDI (Figure 2C).

FIGURE 2.

Preparation and characterization of ALSF at various GPX4‐siRNA:Fe^{3 +} mass ratios. (A) Particle size distribution, (B) particle size and PDI immediately after mixing. (C) Particle size and PDI after formulation. (D)TEM image of ALSF. Scale bar: 100 nm. (E) Schematic representation of ALSF morphology. (F) Zeta potential after formulation. (G) Agarose gel electrophoresis and (H) encapsulation efficiency of GPX4‐siRNA. (I) Encapsulation efficiency of Fe^{3 +}. (J) Changes in particle size during incubation in 10% FBS at 37°C. All experiments were repeated three times (n = 3). ns, not significant.

FIGURE 1.

The mechanism diagram of ALSF selectively inducing ferroptosis in cancer cells.

Subsequently, we performed Transmission Electron Microscopy (TEM) characterization on the ALSF at various GPX4‐siRNA:Fe3⁺ mass ratios. All nanoparticles exhibited uniform spherical shapes, yet subtle differences were observed in their internal architectures (Figure 2D). Integrating the above dynamic light scattering (DLS) results, we illustrated the self‐assembly pathway of ALSF with a schematic diagram (Figure 2E). According to TEM observations, the core‐shell structure was exclusively absent in the ALNP group without GPX4‐siRNA. This phenomenon indicates that the nanoparticle inner core corresponds to the nucleic acid encapsulation domain. As a classical fatty acid, AA triggers electrostatic repulsion either between AA molecules or between AA and nucleic acids during ALNP and ALS assembly. The generated electrostatic repulsion obstructed self‐assembly, further causing poor monodispersity of resultant nanoparticles. Moreover, a prominent bimodal distribution appeared in the ALS group, suggestive of unsuccessful nucleic acid encapsulation in a large proportion of ALS nanoparticles (Figure 2A). TEM images further corroborated the existence of empty‐shell ALS nanoparticles (Figure S2). Although purified ALNP and ALS possessed regular morphology, their defective self‐assembly behavior during mixing could adversely affect nanoparticle yield, empty‐shell fraction, and batch‐to‐batch reproducibility. For Fe³⁺‐doped ALSF nanoparticles, increasing Fe³⁺ dosage enlarged the overall particle size and inner core volume, and simultaneously improved nanoparticle homogeneity. Notably, ALSF (1:0.66 group) showed the optimal performance. Nevertheless, excessive Fe³⁺ led to serious aggregation of the inner core of ALSF (1:1 group) nanoparticles. This indicates that Fe³⁺ enhances self‐assembly control, reduces size heterogeneity, and improves formulation stability. These results highlight the importance of controlling Fe³⁺ concentration.

All ALSF formulations exhibited slightly negative surface charges (Figure 2F). No significant differences were observed in siRNA encapsulation efficiency (Figure 2G, H). All ALSF formulations effectively encapsulated Fe^{3 +}, achieving ∼90% efficiency (Figure 2I). Colloidal stability is critical for in vivo delivery [56, 57]. All ALSF formulations remained stable at 4°C over 7 days, with no significant changes in size or PDI (Figures S3 and S4). However, in Tris‐HCl buffer with 10% fetal bovine serum (FBS, pH 7.4, 37°C), Fe^{3 +} significantly influenced nanoparticle stability. ALNP, ALS, and ALSF with low Fe^{3 +} (1:0.1) rapidly grew from 80 nm to over 200 nm within 6 h. The 1:0.33 group exhibited a slower increase, from 108.6 ± 0.5 nm to 195.8 ± 3.5 nm. Notably, the 1:0.66 group showed optimal stability, with size increasing only from 130.4 ± 1.0 nm to 150.5 ± 1.2 nm over 6 h, likely due to normal serum protein adsorption, and remained stable for 48 h (Figure 2J). Regarding PDI, all groups lacking sufficient Fe^{3 +} increased rapidly above 0.4 within 6 h, indicating structural disruption. In contrast, the 1:0.66 group's PDI rose gradually from 0.223 to 0.278 in 6 h and remained below 0.3 for 30 h (Figure S5). These results demonstrate that ALSF with a 1:0.66 GPX4‐siRNA:Fe³⁺ ratio possesses excellent colloidal stability. It effectively resists serum protein adsorption and ionic interference, providing a robust foundation for prolonged circulation and targeted in vivo delivery.

2.2. ALSF Efficiently Induces Ferroptosis in Ocular Cancer Cells

To determine the optimal Fe^{3 +} ratio in ALSF, we first assessed the silencing efficiency of GPX4 mRNA in C918 cancer cells. As shown in Figure 3A, all ALSF formulations containing 2 µg/mL GPX4‐siRNA achieved approximately 90% knockdown of GPX4 mRNA compared to the ALNP, with no significant differences among groups. At a lower concentration (0.5 µg/mL), however, the ALS and 1:0.1 groups showed significantly reduced knockdown efficiency compared to the 1:0.33 and 1:0.66 groups. These results indicate that Fe^{3 +} contributes substantially to siRNA delivery efficiency.

FIGURE 3.

Mechanistic study of ALSF‐induced ferroptosis in ocular cancer cells. (A) GPX4 mRNA knockdown efficiency of ALSF at different GPX4‐siRNA:Fe^{3 +} mass ratios. (B) Cytotoxicity of ALSF against C918 cancer cells at varying mass ratios. ROS levels in C918 cells following ALSF treatment, shown as fluorescence images (C) and flow cytometry histograms (D). Scale bar: 30 µm. GPX4 protein expression in C918 cells analyzed by western blot (E) and quantification (F). (G) LPO product MDA detection in vitro. LPO in C918 cells after ALSF treatment, shown as fluorescence images (H) and flow cytometry histograms (I). Scale bar: 30 µm. (J) TEM images of C918 cells following ALSF treatment. Scale bar: 500 nm. All experiments were repeated three times (n = 3). ns, not significant, ^* p < 0.1, ^** p < 0.01, and ^**** p < 0.0001.

Next, we evaluated the cytotoxic effects of these formulations on C918 cancer cells and 293T normal cells to identify the optimal Fe^{3 +} ratio. As shown in Figure 3B and Figure S6, ALNP and ALS showed no significant cytotoxicity across all tested concentrations (0.25–2 µg/mL), confirming the biocompatibility of the carrier. Although ALS effectively silenced GPX4 mRNA, it did not induce notable cell death, likely due to the co‐delivered AA, which may exert antioxidant effects under normal oxidative conditions, partially counteracting the pro‐oxidant effect of GPX4 silencing [25, 58, 59]. In contrast, ALSF formulations containing Fe^{3 +} (1:0.1, 1:0.33, 1:0.66) demonstrated potent and selective cytotoxicity against C918 cells, reducing viability to 20–30%, while 293T cells remained >80% viable at most concentrations. The only exception was the 1:0.1 group at low concentrations (0.25–0.5 µg/mL), which showed minimal killing. Notably, ALSF exhibited a pharmacodynamic threshold for cancer cell killing. For the 1:0.1 group, minimal cytotoxicity occurred at 0.25–0.5 µg/mL, whereas viability sharply dropped to ∼80% at 1 µg/mL. For the 1:0.33 and 1:0.66 groups, significant cytotoxicity (∼80%) was observed at 0.5 µg/mL or higher.

This threshold‐dependent effect likely arises from the synergistic modulation by ALSF of three ferroptosis drivers: GPX4 activity, intracellular iron, and LPO accumulation. At low doses, perturbations are insufficient to disrupt redox homeostasis. Once the threshold is exceeded, these factors synergize to rapidly disrupt redox balance, triggering cancer cell death [60, 61]. These results also highlight the differential sensitivity of cancer versus normal cells, consistent with ferroptosis’ metabolic basis. Cancer cells’ distinct redox and iron metabolism render them more susceptible to iron‐dependent LPO.

Based on these findings, the 1:0.33 and 1:0.66 formulations required lower effective doses than the 1:0.1 group, with no significant difference between them. Considering additional characterization data (Figure 2), the 1:0.66 formulation demonstrated superior stability, efficacy, and safety. Therefore, GPX4‐siRNA:Fe^{3 +} = 1:0.66 was used in all subsequent experiments.

As illustrated in the schematic of ALSF‐induced ferroptosis (Figure 1), ALSF enters cancer cells via endocytosis and localizes to lysosomes, where Fe^{3 +} and AA are released. In the acidic lysosomal environment, iron recycling occurs, replenishing intracellular iron and generating Fe^{2 +}. Fe^{2 +} subsequently reacts with H₂O₂ via the Fenton reaction, producing reactive oxygen species (ROS). We measured ROS levels in C918 cells to investigate this mechanism. ROS levels in the ALNP were similar to controls. In the ALS group, GPX4 protein knockdown impaired ROS scavenging, causing a modest ROS increase. In contrast, ALSF generated substantial ROS due to Fe^{3 +} delivery and iron recycling, combined with GPX4 silencing, resulting in significantly higher intracellular ROS than other groups (Figure 3C,D).

Western blot analysis confirmed GPX4 protein knockdown by ALSF (Figure 3E,F). Raw Western blot images are shown in the supplementary data (Figure S7). At high doses, both ALS and ALSF reduced GPX4 protein by ∼60%, with no significant difference. At low doses, however, ALSF containing Fe^{3 +} still achieved significant knockdown, whereas ALS alone did not.

Inspired by the industrial electro‑Fenton technology characterized by electrochemical iron cycling, Liu et al. constructed a nanoengineered system based on Fe³⁺/tannic acid (TA) coordination, which achieves iron redox cycling via chemical reduction of Fe³⁺ to Fe²⁺ by TA, thereby inducing ferroptosis [21]. In contrast to this iron‑centric strategy, our study employs AA as a multifunctional component in the ALNP system, serving three roles: a structural lipid, a reducing agent for Fe³⁺, and—most critically—a substrate for LPO.

Among these, the role of AA as an LPO substrate is of particular significance, as the availability of PUFA‑containing phospholipid substrates has been identified as the primary rate‑limiting factor in ferroptosis‑based cancer therapy. Indeed, multiple studies have demonstrated that PUFA deficiency confers resistance to ferroptosis, while exogenous supplementation with PUFAs (including AA) can effectively sensitize cancer cells to ferroptosis [62, 63, 64, 65, 66, 67]. To validate this rationale, we directly confirmed in vitro that AA can act as an exogenous substrate for LPO, thereby increasing LPO accumulation and elevating the production of malondialdehyde (MDA), a terminal product of lipid peroxidation (Figure 3G). Next, according to the proposed mechanism, ALSF promotes ROS generation via iron recycling and the Fenton reaction while inhibiting GPX4, leading to LPO accumulation. LPO levels measured in C918 cells confirmed this effect: the ALSF group showed significantly higher LPO than the control, empty ALNP, or ALS groups (Figure 3H,I), indicating that iron‐dependent AA recycling drives LPO accumulation. Furthermore, TEM revealed characteristic ultrastructural features of ferroptosis in ALSF‑treated C918 cells (Figure 3J), including shrunken mitochondria with reduced or absent cristae, increased matrix density, vesicular protrusions on the plasma membrane with disorganized lipid bilayers, and enlarged lysosomes with disrupted membranes. Collectively, these findings confirm that ALSF efficiently induces ferroptosis in cancer cells.

The multifunctionality of AA—as a structural lipid, a Fe³⁺ reductant, and a LPO substrate—well embodies the design philosophy of “materials as therapeutics,” where a single component simultaneously contributes to delivery, activation, and execution of therapeutic effect. Moreover, ALSF is constructed on the clinically validated LNP platform, which offers a simple preparation process, high reproducibility, and established industrial scalability, with several LNP‑based products already approved for clinical use. This system not only achieves potent ferroptosis‑inducing efficacy against cancer cells but also holds considerable promise for translational development.

2.3. Acid‐Responsive Release, Iron Cycling and Lysosomal Escape Mechanism of ALSF

To investigate the mechanism of Fe^{3 +} and siRNA release by ALSF and its promotion of lysosomal escape in vivo, we designed a series of experiments simulating the physiological environment. Based on reports that phosphate and carboxyl groups undergo protonation under acidic conditions [68], we hypothesized that in lysosomes, phosphate groups of nucleic acids and carboxyl groups of AA are protonated. This alters ALSF structure, facilitating Fe^{3 +} release (Figure 4A). Our experiments showed (Figure 4B) that Fe^{3 +} release was minimal at pH 7.4, but reached ∼90% within 24 h at pH 5.0, simulating the lysosomal environment. Upon prolonged exposure to an acidic environment, SM‐102 in ALSF undergoes a phase transition to an inverted hexagonal structure, which facilitates rapid siRNA release from the nanoparticles [69, 70]. Concurrently, the removal of positively charged Fe³⁺ from ALSF enhances the electrostatic repulsion between AA and nucleic acids, potentially reducing nanoparticle stability and accelerating siRNA release—a phenomenon supported by our previous formulation experiments (Figure 2). As a result, the cumulative siRNA release reached 51% after 72 h of incubation at pH 5.0, whereas under pH 7.4 conditions, the release remained below 20%. After 72 h at pH 5.0, 51% of siRNA was released, compared to <20% at pH 7.4.

FIGURE 4.

Acid‐responsive release, iron cycling, and lysosomal escape of ALSF. (A) Schematic illustration of the ALSF iron cycling mechanism. (B) Release profiles of ALSF under different pH conditions. (C) In vitro demonstration of iron cycling. (D) Cellular validation of iron cycling. (E) Fluorescence images of ROS in C918 cells. Scale bar: 30 µm. (F) Fluorescence images of C918 cells after 4 h incubation with CY5‐labeled ALSF (red); lysosomes labeled with Lysotracker (green). Scale bar: 10 µm. (G) Pearson's correlation coefficient for co‐localization analysis calculated using ImageJ. All experiments were repeated three times (n = 3). ns, not significant; ^*** p < 0.001.

As an easily oxidizable PUFA, AA can initiate iron‐dependent lipid peroxidation. Its oxidation generates lipid radicals, peroxyl radicals, and lipid hydroperoxides, triggering a chain reaction while simultaneously reducing Fe³⁺ to Fe²⁺, thereby sustaining Fenton reactions and elevating ROS levels [71, 72, 73, 74]. Because Fe^{2 +} is much more reactive than Fe^{3 +}, this iron cycle continuously supplies Fe^{2 +}, accelerating ferroptosis. The mechanism is illustrated in Figure 4C. Interestingly, we found that this process is accelerated under acidic conditions both in vitro and at the cellular level, which may be attributed to the fact that the lipid peroxidation chain reaction initiated by AA and Fe³⁺ proceeds more readily in an acidic environment [75, 76, 77, 78, 79].To confirm AA‐mediated iron cycling under acidic conditions, we used o‐phenanthroline, which forms an orange complex with Fe^{2 +} but not Fe^{3 +}. At pH 7.4, Fe^{3 +} reduction was minimal after 6 h. At pH 5.0, AA effectively reduced Fe^{3 +} to Fe^{2 +}, which could be oxidized back to Fe^{3 +} by H₂O₂ (Figure 4D). Achieving comparable Fe^{2 +} generation at pH 7.4 required 48 h, highlighting the role of acidic protonation in Fe^{3 +} dissociation and redox cycling (Figure S8).

We next validated iron cycling at the cellular level. Fe^{2 +} reacts with H₂O₂ to produce ROS; thus, intracellular ROS indirectly reflects Fe^{2 +} production. ROS assays confirmed the in vitro findings: ALSF significantly increased ROS in C918 cells. Pre‐treatment with the iron chelator DFO prevented ROS elevation, even with concurrent GPX4 knockdown, demonstrating that iron ions drive the ferroptosis process (Figure 4E). To further probe the role of acidity, we delayed lysosomal acidification with ammonium chloride, which markedly reduced ALSF‐induced ROS. Buffering culture media to pH 5.0 enhanced ROS compared to pH 7.4, confirming the dependence of ALSF‐induced ROS on acidic conditions.

LNP, as a class of nanocarriers, are primarily internalized by mammalian cells via clathrin‐mediated endocytosis and macropinocytosis, following which they are sequestered in early endosomes and progressively mature into late endosomes, ultimately fusing with degradative lysosomes. To achieve functional cytosolic delivery of the encapsulated nucleic acid cargo, LNPs must destabilize the endosomal membrane and release their payload into the cytosol before lysosomal degradation occurs—a process termed endosomal (or lysosomal) escape, which typically takes place during the transition from early to late endosomes. This escape is predominantly mediated by the ionizable lipid component: upon protonation in the acidic endosomal environment (pH ≈ 5.5–6.5), the ionizable lipids acquire a positive charge and interact with anionic endosomal phospholipids, inducing membrane fusion, formation of inverted hexagonal (HII) phases, and transient membrane disruption, thereby enabling cargo release into the cytosol. Despite its critical importance, endosomal escape remains a major rate‐limiting step, with only approximately 1–2% of internalized LNPs successfully delivering their cargo to the cytosol [80, 81, 82, 83].

To track the intracellular fate of ALSF and its lysosomal retention, we encapsulated Cy5‐labeled nucleic acids into ALSF and incubated them with cells. Lysosomes were stained with Lysotracker Green at 1, 2, 4 and 12 h post‐uptake. Quantitative colocalization analysis showed high Pearson's correlation (R = 0.89±0.04) between ALSF and lysosomes at early time points (0–1 h) (Figures S9 and S10). The colocalization decreased progressively thereafter, suggesting that the nanoparticles escaped from lysosomes. Fluorescence images (Figure 4F) showed that both ALS and ALSF facilitated nucleic acid escape from lysosomes at 4 h post‐treatment. However, ALSF exhibited significantly higher escape efficiency in C918 cells than ALS, as reflected by a lower Pearson correlation coefficient. In normal cells, escape efficiency was lower (Figure 4G). The enhanced escape by ALSF likely arises from two mechanisms. First, lysosomal protons compete with Fe^{3 +} for binding to AA carboxyl and nucleic acid phosphate groups, enhancing the proton sponge effect [84]. Second, ROS generated by iron cycling can disrupt lysosomal membranes, facilitating nucleic acid delivery [85, 86, 87]. Cancer cells, which have higher H₂O₂ levels, produce more ROS than normal cells, resulting in selective nucleic acid delivery and ferroptosis at lower doses, ensuring safety in normal tissues. This experimental observation provides a clear explanation for why ALSF can efficiently knock down GPX4 mRNA and protein in cancer cells (Figure 3A,E) at low doses and exhibit selective cytotoxicity toward cancer cells (Figure 3B and Figure S6).

2.4. Selective Cytotoxicity of ALSF in Cancer Cells Is Driven by Elevated H₂O₂ Levels

To confirm that the selective cytotoxicity of ALSF arises from higher H₂O₂ levels in cancer cells, we evaluated three cancer cell lines and three normal cell lines in vitro. ALSF induced over 50% cytotoxicity in cancer cells, including C918, non‐small cell lung cancer (A549), and pancreatic cancer (PANC‐1) (Figure 5A). In contrast, it showed minimal toxicity in normal cells, including mouse fibroblasts (3T3), human renal epithelial cells (293T), and human hepatic stellate cells (LX2) (Figure 5B). This selectivity is attributed to the substantially lower H₂O₂ levels in normal cells [88].

FIGURE 5.

Differential cytotoxicity of ALSF in normal and cancer cells. (A) Cytotoxicity of ALSF in various cancer cell lines. (B) Cytotoxicity of ALSF in various normal cell lines. (C) Fluorescence images of ROS (green) and Fe^{2 +} (red) in cells treated with ALSF. Scale bar: 170 µm. All experiments were repeated three times (n = 3). ns, not significant, ^* p < 0.01 and ^**** p < 0.0001.

Upon addition of 10 µM H₂O₂, ALSF cytotoxicity in cancer cells remained largely unchanged, whereas toxicity in normal cells increased markedly, reaching levels comparable to those in cancer cells (Figure 5B). Fluorescence imaging further confirmed that normal cells treated with ALSF plus H₂O₂ exhibited ROS levels and Fe^{2 +} content similar to cancer cells treated with ALSF alone (Figure 5C).

Together, these results demonstrate that ALSF‐mediated ferroptosis selectively targets cancer cells due to their elevated H₂O₂ levels, while sparing normal cells under physiological conditions.

2.5. In Vivo Antitumor Efficacy and Biosafety Evaluation of ALSF

The promising in vitro results prompted us to further evaluate the in vivo tumor inhibition and biosafety of ALSF. A xenograft model was established by subcutaneously injecting C918 cells into female BALB/c mice. Experiments began when tumor volumes reached approximately 100 mm³ (Figure 6A). First, ALSF encapsulating CY5.5‐labeled nucleic acids was administered via tail vein injection. After 48 h, tumors and major organs were collected and imaged using an in vivo fluorescence system. It has been reported that upon systemic administration, LNPs adsorb plasma apolipoprotein E (ApoE) on their surface to form a protein corona, which facilitates their penetration through the hepatic sinusoidal fenestrae into the liver parenchyma. Hepatocytes, which highly express the low‐density lipoprotein receptor (LDLR), specifically recognize ApoE, initiating receptor‐mediated endocytosis to internalize the LNPs. Following endosomal escape and cargo release, the ApoE‐mediated lipoprotein metabolism pathway enables efficient hepatic enrichment of LNPs [89, 90]. Given that ApoE adsorption is mediated by the properties of ionizable lipids [90, 91, 92], we observed a similar hepatic accumulation phenomenon in the ALSF group in the in vivo imaging at 24 h post‐injection. However, after 48 h of metabolic clearance in mice, the hepatic signal gradually diminished. Upon dissection and simultaneous imaging of the tumors and visceral organs, we were surprised to find that ALSF had effectively delivered siRNA to the tumor site (Figure 6B). Quantitatively, the fluorescence intensity at tumor sites was markedly higher than that in the liver and kidneys (Figure S11).

FIGURE 6.

In vivo antitumor efficacy and biosafety of ALSF. (A) Treatment regimen schematic in C918 tumor‐bearing mice. (B) Ex vivo fluorescence imaging of tumors and major organs 48 h after CY5‐labeled ALSF injection. (C)Tumor growth curves and (D) representative images of excised tumors on day 18. (E) Body weight changes during the 18‐day treatment period (n = 5). Representative H&E‐stained sections of (F) tumors and (G) livers collected on day 18. Scale bar: 100 µm.

This tumor accumulation is likely attributable to the enhanced permeability and retention (EPR) effect, a core pathophysiological basis of passive targeting for nanomedicines in solid tumors. The EPR effect arises from the unique vascular and lymphatic abnormalities within the tumor microenvironment: tumor neovasculature exhibits disorganized endothelial cell arrangement and discontinuous basement membranes, forming endothelial gaps of tens to hundreds of nanometers, through which circulating nanocarriers can extravasate into the tumor interstitium. Concurrently, the underdeveloped and dysfunctional lymphatic drainage within solid tumors fails to efficiently clear macromolecules and nanoparticles from the interstitium, resulting in selective retention and prolonged accumulation of nanocarriers in tumor tissues [93, 94, 95]. Furthermore, as previously mentioned, the siRNA release from ALSF is considerably accelerated under acidic conditions compared to normal physiological environments. Taken together, the synergistic effect of the passive targeting (EPR effect) and the acid‐responsive release property of ALSF likely accounts for its preferential tumor accumulation.

Next, we assessed in vivo tumor inhibition. Twenty‐five tumor‐bearing mice were randomly divided into five groups and monitored for 18 days. The empty ALNP group received the carrier at a dosage equivalent to that of ALSF (based on GPX4‐siRNA content, 0.7 mg/kg). Each group received intravenous injections every three days for a total of five doses (Figure 6A). PBS served as a negative control. Compared to the PBS and ALNP groups, ALSF treatment significantly suppressed tumor growth (Figure 6C,D). At 0.35 mg/kg, ALSF achieved approximately 40% tumor inhibition, comparable to the ALS group at 0.7 mg/kg. At 0.7 mg/kg, ALSF achieved up to 75% tumor inhibition.

H&E staining of tumor tissues further confirmed these findings (Figure 6F). Tumors from PBS and ALNP groups retained mostly intact cell morphology. In contrast, ALS and ALSF treatment groups showed extensive loss of cellular integrity, with ALSF at 0.7 mg/kg producing the most pronounced disruption. The tumor cell killing effect was comparatively weaker in the ALS (0.7 mg/kg) and ALSF (0.35 mg/kg) groups. Overall, the in vivo antitumor efficacy mirrored the in vitro cytotoxicity results.

No significant differences in body weight were observed among the groups (Figure 6E). H&E staining of liver tissues collected at the end of the experiment revealed no apparent pathological abnormalities in any treatment group (Figure 6G), indicating good biosafety of ALSF.

3. Conclusion

In this study, we developed a novel iron‐coordinated LNP, ALSF, for selective induction of ferroptosis in cancer cells. ALSF efficiently co‐delivers AA, Fe^{3 +}, and GPX4‐siRNA, enabling synergistic regulation of three key ferroptosis drivers: GPX4 activity, intracellular iron, and LPO. We demonstrated that the introduction of Fe^{3 +} enhances nanoparticle stability, controls siRNA release, and promotes lysosomal escape via both proton sponge effect and ROS‐mediated membrane disruption. This dual functionality allows ALSF to achieve high siRNA delivery efficiency selectively in cancer cells. In vitro, ALSF induced potent, threshold‐dependent cytotoxicity in multiple cancer cell lines while sparing normal cells. This selectivity is attributed to higher H₂O₂ levels in cancer cells, which drive iron cycling and ROS generation. Mechanistic studies confirmed that ALSF triggers substantial LPO and morphological features consistent with ferroptosis. In vivo, ALSF accumulated efficiently in tumor tissue, suppressed tumor growth in a dose‐dependent manner, and exhibited excellent biosafety. Notably, the antitumor efficacy of ALSF at half the siRNA dose was superior to ALS lacking Fe^{3 +}, highlighting the synergistic effect of iron coordination. Overall, our findings present a rational design of multifunctional nanoparticles that exploit tumor‐specific redox and iron metabolism. This strategy provides a new paradigm for selective ferroptosis induction and targeted cancer therapy, combining high efficacy, selectivity, and safety.

4. Experimental Section

4.1. LNP Synthesis and Characterization

ALNP and ALS were prepared using microfluidic mixing, following established LNP protocols [48]. For ALNP, AA was used to replace DSPC as the structural lipid. AA, SM‐102, cholesterol, and DMG‐PEG2000 (all purchased from SINOPEG) were dissolved in ethanol at defined molar ratios to form the lipid phase. For ALNP, 20 mM Tris‐HCl buffer (pH 5.0, AVT) served as the aqueous phase. For ALS, siRNA (Promega) was dissolved in the same aqueous phase. The aqueous and lipid phases were rapidly mixed at a 3:1 volume ratio (aqueous:lipid) using a microfluidic system (INanoL, Maianna, China) with an N/P ratio of 6. The resulting solution was immediately diluted threefold with 20 mM Tris‐HCl (pH 7.4) and concentrated in an ultrafiltration tube (MWCO 100 kDa, Millipore) to one‐third of the original volume. This dilution–concentration cycle was repeated three times to remove ethanol and replace the buffer. The final product was concentrated to the desired concentration to obtain ALNP or ALS. For ALSF, FeCl₃·6H₂O (Sinopharm, Fe^{3 +}/siRNA, w/w) was added to the lipid ethanol phase at specified mass ratios before mixing; all subsequent steps were identical to ALS preparation. Particle size, PDI, and zeta potential were measured using DLS (Zetasizer Nano ZS). Encapsulation efficiencies of siRNA and Fe^{3 +} were determined by agarose gel electrophoresis and inductively coupled plasma–mass spectrometry (ICP‐MS, NexION 300X), respectively. Grayscale values were analyzed using ImageJ software.

4.2. Cell Culture

The human uveal melanoma cell line C918 (choroid origin, Procell Life Science & Technology Co., Ltd., cat. no. CL‐0264) was cultured in RPMI 1640 medium (Thermo Fisher Scientific, Inc., cat. no. C11875500BT) supplemented with 10% fetal bovine serum (FBS; Vivacell, cat. no. C04001‐500) and 1% penicillin–streptomycin solution (KeyGEN BioTECH, cat. no. KGL2303‐100) at 37 °C under 5% CO₂. The human pancreatic ductal adenocarcinoma cell line Panc‐1 and hepatic stellate cell line LX‐1 were obtained from Zhejiang Meisen Cell Technology Co., Ltd. The human non‐small cell lung cancer cell line A549, human embryonic kidney cell line 293T, and murine embryonic fibroblast cell line 3T3 were purchased from Hunan Fenghui Biotechnology Co., Ltd. These cell lines were cultured in Dulbecco's Modified Eagle Medium (DMEM, Gibco, cat. no. C11995500BT) containing 10% FBS and 1% penicillin–streptomycin under the same conditions (37 °C, 5% CO₂).

4.3. Quantitative Real‐Time PCR (q‐PCR)

C918 cells were seeded in 6‐well plates at 3 × 10⁵ cells per well and cultured until ∼80% confluence. Cells were then treated with PBS, ALNP, or ALSF at 3 µg/mL for 24 h. Total RNA was extracted using Trizol reagent according to the manufacturer's protocol, and RNA concentration and purity were measured with a Nanodrop 2000 spectrophotometer. Complementary DNA (cDNA) was synthesized from 500 ng of total RNA using the Hifair III 1st Strand cDNA Synthesis SuperMix for qPCR (gDNA digester plus; YEASEN, cat. no. 11141ES60) following the manufacturer's instructions. Quantitative PCR was performed in 20 µL reactions containing 10 µL Hieff UNICON Universal Blue qPCR SYBR Green Master Mix (YEASEN, cat. no. 11184ES08), 2 µL gene‐specific primers, and 2 µL diluted cDNA. Primer pairs used were: human GPX4 (Beyotime Biotechnology, cat. no. QH00012S) and human β‐actin (ACTB; Beyotime Biotechnology, cat. no. QH00002S). Thermal cycling conditions were 95 °C for 2 min, followed by 40 cycles of 95 °C for 10 s and 60 °C for 30 s. Relative gene expression was calculated using the 2^‐ΔΔCt method, with β‐actin as the internal reference.

4.4. Cell Viability

Six cell lines were used: normal cells (293T human embryonic kidney, LX2 human hepatic stellate, and 3T3 mouse embryonic fibroblasts) and cancer cells (C918 human uveal melanoma, PANC‐1 human pancreatic cancer, and A549 human lung cancer). Cells were seeded in 96‐well plates at 5 × 10³ cells per well and cultured overnight for attachment. Experimental groups included blank (medium only), control (untreated), material control (ALNP), and experimental (ALSF), all at a fixed dose of 2 µg/mL. Selected wells were additionally treated with hydrogen peroxide to induce oxidative stress. After 24 h treatment, 10 µL of CCK‐8 reagent was added per well and incubated at 37 °C for 1–4 h. Absorbance was measured at 450 nm using a microplate reader, and cell viability was calculated accordingly.

4.5. Intracellular ROS and LPO Assays

C918 cells were seeded in 6‐well plates and cultured overnight at 37 °C in a 5% CO₂ incubator. After attachment, cells were assigned to six groups: blank control, negative control, ALNP (2 µg/mL), ALS (2 µg/mL), ALSF (2 µg/mL), and positive control (Rosup, 1:1000 dilution added 20–30 min before staining), with three replicates per group. After 24 h treatment, the medium was removed, and cells were washed twice with PBS. ROS Measurement: Intracellular ROS was detected using the Reactive Oxygen Species Assay Kit (DCFH‐DA; Beyotime, cat. no. S0033S) following the manufacturer's instructions. Cells were incubated with 1 mL of DCFH‐DA working solution (10 µM, diluted 1:2000 in serum‐free medium) at 37 °C for 15 min in the dark. For fluorescence imaging, cells were washed, 2 mL PBS was added, and images were immediately captured. For flow cytometry, cells were harvested by trypsinization, washed twice with PBS, resuspended in 200–300 µL PBS, and analyzed on a flow cytometer (FL1 channel, 1 × 10⁴ cells per sample). Mean fluorescence intensity was quantified using FlowJo software. LPO Measurement: LPO was assessed using the Lipid Peroxidation Assay Kit (BODIPY 581/591 C11; Beyotime, cat. no. S0043S). Cells were incubated with 1 mL of staining solution (2 µM in serum‐free medium) at 37 °C for 20 min in the dark. For fluorescence imaging, 2 mL PBS was added, and oxidized green fluorescence (Ex/Em = 488/510 nm) was immediately captured. For flow cytometry, cells were harvested, washed twice with PBS, resuspended in 200–300 µL PBS, and analyzed (FL1 channel, 1 × 10⁴ cells per sample). LPO levels were quantified using FlowJo software.

4.6. Lipid Peroxidation (MDA) Assay

The level of lipid peroxidation was evaluated by measuring malondialdehyde (MDA) using the thiobarbituric acid (TBA) method. The following treatment groups were established: control (UP water), AA (200 µM), H₂O₂ (100 µM), Fe^{2 +} (100 µM), AA (200 µM) + H₂O₂ (100 µM), AA (200 µM) + H₂O₂ (100 µM) + Fe^{2 +} (100 µM), ALSF (50 µg/mL), ALSF (50 µg/mL) + H₂O₂ (100 µM), and AA (200 µM) + ALSF (50 µg/mL) + H₂O₂ (100 µM). All samples were incubated at 37°C for 6 h in a final reaction volume of 0.1 mL. According to the manufacturer's instructions for the Lipid Peroxidation (MDA) Assay Kit (cat. no. S0131S; Beyotime), 0.1 mL of each sample, standard, or blank was mixed with 0.2 mL of MDA working solution. The mixtures were heated in a boiling water bath at 100°C for 15 min (with proper sealing to prevent boiling over), then cooled to room temperature. After centrifugation at 1000 × g for 10 min, 200 µL of the supernatant was transferred to a 96‐well plate, and the absorbance was measured at 532 nm using a microplate reader. The MDA concentration in each sample was calculated based on a standard curve.

4.7. Western Blotting Assay

C918 cells were seeded in 6‐well plates and cultured overnight. Cells were treated with PBS, ALNP, ALS, or ALSF for 24 h. Total protein was extracted using the Membrane, Nuclear and Cytoplasmic Protein Extraction Kit (KeyGEN BioTECH, cat. no. KGB5300‐50), and protein concentrations were measured with the BCA Protein Assay Kit (KeyGEN BioTECH, cat. no. KGB2101‐250). Equal amounts of protein (20 µg) were separated on 12% SDS‐PAGE and transferred to 0.45 µm PVDF membranes (Biosharp, cat. no. BS‐PVDF‐45). Membranes were blocked with 5% non‐fat milk for 1 h at room temperature, then incubated overnight at 4 °C with primary antibodies against GPX4 (Rabbit pAb, Servicebio, cat. no. GB115275) and β‐actin (Rabbit pAb, Servicebio, cat. no. GB15003). Membranes were subsequently incubated with HRP‐conjugated goat anti‐rabbit IgG (H+L) (CUSABIO, cat. no. CSB‐PA000106RA01HU) for 1.5 h at room temperature. Protein bands were visualized using the ECL Chemiluminescent Substrate Kit (Super Sensitive, Biosharp, cat. no. BL520B) and imaged with a chemiluminescence system. Band intensities were quantified using ImageJ software, with β‐actin serving as the loading control.

4.8. Transmission Electron Microscopy (TEM)

The morphology of the nanoparticles was characterized by TEM using the negative staining method. ALNP, ALS, and ALSF samples were diluted with ultrapure water to a final lipid concentration of 0.1–1.0 mg/mL. A clean copper grid was placed on a piece of filter paper, and 30 µL of the diluted sample was gently dropped onto the grid and allowed to stand at room temperature for approximately 10 min to ensure sufficient sample adsorption. Excess liquid was carefully blotted away with filter paper. Subsequently, 30 µL of phosphotungstic acid (PTA) staining solution was added to the grid and incubated for 2 min at room temperature for negative staining. The excess dye was removed by blotting with filter paper, and the grid was then dried under an infrared lamp. The prepared samples were observed and photographed using a transmission electron microscope (JEM‐2100Plus; JEOL Ltd., Tokyo, Japan).

C918 cells were seeded in 6‐well plates at an appropriate density. After attachment, cells were treated with ALSF (2 µg/mL). At 48 h post‐treatment, when cells reached 70%–80% confluence, they were collected using a cell scraper and transferred into 1.5 mL centrifuge tubes. Cells were centrifuged to form a visible pellet (∼size of a mung bean). The supernatant was carefully removed, and the pellet was gently overlaid with pre‐cooled 2.5% glutaraldehyde fixative (for electron microscopy; Biosharp, cat. no. BL911A) along the tube wall without disturbing the pellet. Tubes were sealed with Parafilm, left at room temperature for 30 min, and then stored at 4 °C. Samples were subsequently sent to the Shiyanjia platform for TEM imaging.

4.9. In Vitro Acid‐Responsive Release Assay

To assess the release profiles of siRNA and Fe^{3 +} under different pH conditions, ALSF (CY5‐siRNA, 100 µg/mL) was placed in a dialysis cassette (MWCO 20 kDa) and immersed in 100 volumes of buffer at either pH 7.4 or pH 5.0 (20 mM Tris‐HCl) to simulate physiological and lysosomal environments, respectively. The system was incubated at 25 °C under magnetic stirring (300 rpm). At predefined time points, aliquots of the ALSF solution remaining inside the dialysis cassette were collected. Fe^{3 +} concentrations were measured using inductively coupled plasma optical emission spectroscopy (ICP‐OES), while CY5.5‐siRNA release was quantified by UV–vis spectrophotometry.

4.10. In Vitro Validation of Iron Cycling

This experiment aimed to visually demonstrate Fe^{3 +}/Fe^{3 +} generation and validate the iron cycling process in vitro using the differential chromogenic reaction of Fe^{3 +}/Fe^{2 +} with 1,10‐phenanthroline. Fe^{2 +} rapidly forms a stable orange‐red complex with 1,10‐phenanthroline at pH 3–9, whereas Fe^{3 +} does not react. The following groups were prepared: Fe^{3 +} alone, Fe^{3 +} alone, Fe^{3 +} + AA at pH 7.4, Fe^{3 +} + AA at pH 5.0, and Fe^{2 +} + H₂O₂. Each sample was incubated at 37 °C for 6 h. After incubation, an equal volume of 1,10‐phenanthroline solution (1 mg/mL) was added. Samples were allowed to stand for 5 min to stabilize color, after which the reactions were observed and photographed.

4.11. Cellular Validation of Iron Cycling

C918 cells were seeded in 6‐well plates and cultured overnight at 37 °C in a 5% CO₂ incubator. Cells were assigned to the following treatment groups: control, ALSF + DFO, ALSF + NH₄Cl, ALSF at pH 7.4, ALSF and ALSF at pH 5.0.

ALSF + DFO group: Cells were treated with 100 µM DFO and 2 µg/mL ALSF for 24 h.

ALSF + NH₄Cl group: Cells were treated with 30 mM NH₄Cl and 2 µg/mL ALSF for 24 h.

ALSF pH 7.4 group: 2 µg/mL ALSF was dissolved in culture medium containing 10% v/v 20 mM Tris‐HCl buffer (pH 7.4) and incubated for 24 h. ALSF group: 2 µg/mL ALSF was dissolved in culture medium containing 10% v/v and incubated for 24 h. ALSF pH 5.0 group: 2 µg/mL ALSF was dissolved in culture medium containing 10% v/v 20 mM Tris‐HCl buffer (pH 5.0) and incubated for 24 h. Control group: Received an equal volume of Tris‐HCl buffer.

After treatment, the medium was removed, and cells were washed twice with PBS. Intracellular ROS levels were measured using the Reactive Oxygen Species Assay Kit (cat. no. S0033S; Beyotime) according to the manufacturer's protocol, as described above.

4.12. Co‐localization Analysis of CY5‐Labeled ALSF with Lysosomes

C918 and 293T cells were seeded in 6‐well plates and cultured at 37 °C with 5% CO₂ for 24 h until reaching 70%–80% confluence. Cells were then treated with 3 µg/mL CY5‐labeled ALSF or CY5‐labeled ASF for 4 h. After treatment, the medium was removed, and cells were washed twice with serum‐free medium. A green fluorescent lysosomal probe (diluted to working concentration in serum‐free medium) was added, and cells were incubated at 37 °C for 30 min in the dark. Following incubation, the staining solution was removed, and cells were washed 2–3 times with pre‐warmed PBS. Fluorescence images were captured using an inverted fluorescence microscope (FITC channel for lysosomes, CY5 channel for formulations). Co‐localization was quantified using ImageJ software, and Pearson correlation coefficients were calculated to assess the degree of overlap between formulations and lysosomes, thereby evaluating lysosomal escape efficiency.

4.13. ROS and Fe^{2 +} Double Fluorescence Staining Assay

293T, LX2, 3T3, C918, Panc‐1, and A549 cells were seeded in 24‐well plates and allowed to attach. The treatment groups were: control, H₂O₂ (10 µM for 24 h), ALSF (2 µg/mL for 24 h), and ALSF + H₂O₂ (2 µg/mL ALSF + 10 µM H₂O₂ for 24 h). Staining solutions were prepared according to the Ferroptosis Fluorescent Detection Kit (RhoNox‐6/CM‐H₂DCFDA, cat. no. C11900; Beyotime) instructions by diluting CM‐H₂DCFDA (10 µM) and RhoNox‐6 (5 µM) in serum‐ and phenol red‐free medium. A total of 250 µL of the mixed staining solution was added per well, and cells were incubated at 37 °C for 30 min in the dark. After incubation, the staining solution was removed, and cells were washed with PBS. Fluorescence images were immediately captured using a fluorescence microscope. CM‐H₂DCFDA (green) was detected at Ex/Em = 495/530 nm, and RhoNox‐6 (red) at Ex/Em = 545/585 nm.

4.14. In Vivo Distribution

Five‐week‐old female BALB/c nude mice (Hunan SJA Laboratory Animal Co., Ltd.) were subcutaneously injected with 200 µL of C918 cell suspension (1 × 10⁷ cells) into the right forelimb axilla to establish a tumor xenograft model. When tumor volumes reached ∼300 mm³, mice were intravenously injected via the tail vein with ALSF nanoparticles encapsulating CY5.5‐labeled DNA (0.75 mg/kg). At 48 h post‐injection, mice were euthanized, and tumors and major organs were collected for ex vivo fluorescence imaging to assess the in vivo biodistribution of ALSF.

4.15. In Vivo Pharmacodynamic Study

Five‐week‐old female BALB/c nude mice were obtained from Hunan SJA Laboratory Animal Co., Ltd. (Hunan, China) and maintained under specific pathogen‐free conditions with free access to sterile food and water. C918 cells in logarithmic growth phase were trypsinized, washed twice with sterile PBS, and resuspended at 5 × 10⁷ cells/mL. Each mouse received a subcutaneous injection of 200 µL cell suspension (1 × 10⁷ cells) in the right forelimb axilla. When tumors reached ∼100 mm³, mice were randomly assigned to five groups (n  =  5 per group): (1) control (PBS), (2) ALNP (0.75 mg/kg), (3) ALS (0.75 mg/kg), (4) ALSF low‐dose (0.35 mg/kg), and (5) ALSF high‐dose (0.75 mg/kg). All formulations were administered via tail vein every three days for five doses. Tumor dimensions were measured daily using a digital caliper, and volume was calculated as: volume  =  (length × width²)/2. Body weight was monitored daily to assess systemic toxicity.

On day 18 after the first treatment, mice were euthanized. Tumors were harvested, photographed, and fixed in neutral General Tissue Fixative Solution (cat. no. G1101‐500ML; Servicebio) for histopathology. Liver tissues were also collected for frozen sectioning and H&E staining to evaluate hepatic safety.

Funding

This work was supported by the National Natural Science Foundation of China (Nos. 82170850).

Ethics Statement

All animal experiments were approved by the Ethics Committee of the Xiangya School of Pharmacy, Central South University. (Approval No. CSU‐2023‐XMSB‐1989).

Conflicts of Interest

The authors declare no conflict of interest.

Supporting information

Supporting File: smll74925‐sup‐0001‐SuppMat1.docx.

Data Availability Statement

The data that supports the findings of this study are available in the supplementary material of this article.