Hierarchically Chiral Silver Nanoclusters Mediated Enantioselective Glutathione Depletion and Intracellular Self‐Assembly for Enhanced Anticancer Therapy

ABSTRACT

Chiral nanomedicine design benefits from understanding enantioselective interactions between hierarchically chiral nanoclusters and tumor biomarkers. In this study, we synthesized a pair of silver nanoclusters with bilevel chirality, designated as D‐Ag₆SP₆ and L‐Ag₆SP₆, which were further functionalized with polyethylene glycol (PEG) to enhance their biocompatibility, yielding D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles. Notably, these chiral nanoclusters show enantioselective interactions with glutathione (GSH), as evidenced by their distinct binding constants (K _a). Specifically, the K _a values for L‐Ag₆SP₆ are 2.47 × 10⁷ M⁻¹, while those for D‐Ag₆SP₆ reach 1.33 × 10⁸ M⁻¹, indicating that the D‐enantiomer exhibits significantly stronger GSH‐binding affinity. Both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG deplete intracellular GSH in tumor cells through a ligand‐exchange mechanism, subsequently forming homochiral coordination supramolecular polymers (CSPs) composed of Ag(I)‐GSH complexes. Following adequate aging, these CSPs aggregate into microscale fibers that induce mitochondrial mechanical damage, leading to a significant increase in intracellular reactive oxygen species (ROS) levels, which ultimately induces apoptosis and ferroptosis of tumor cells. Consistently, D‐Ag₆SP₆@PEG demonstrates superior tumor‐killing efficacy in both in vitro and in vivo studies compared with L‐Ag₆SP₆@PEG, attributed to its higher GSH‐binding affinity. This work underscores the significance of chiral design in nanomedicines and provides valuable insights for the development of advanced chiral nanotherapeutics.

Hierarchically chiral silver nanoclusters achieve enantioselective glutathione depletion and initiate intracellular self‐assembly into microfibers. These fibers cause mitochondrial damage that elevates ROS and suppresses GPX4 activity, thereby inducing apoptosis and ferroptosis for efficient tumor inhibition. This work demonstrates the significant potential of chiral nanoclusters in enabling precise cancer therapy through hierarchical chirality‐driven bioeffects.

1. Introduction

Glutathione (GSH), present in millimolar concentrations within mammalian tissues, is a ubiquitous chiral tripeptide that functions as a critical intracellular antioxidant. It plays an essential role in maintaining redox homeostasis and safeguarding normal cells from damage caused by reactive oxygen species (ROS), toxins, and pharmaceutical agents [1, 2, 3]. Notably, cancer cells frequently display elevated levels of GSH, which has spurred the development of therapeutic strategies targeting GSH depletion. This approach holds promise in cancer treatment, as reducing GSH levels can lead to a rapid accumulation of ROS, ultimately inducing apoptotic cell death and ferroptosis [4, 5, 6, 7]. To date, three primary strategies have been developed for GSH depletion in cancer therapy. The first involves direct oxidation of GSH to its oxidized form, glutathione disulfide (GSSG). This method reduces intracellular GSH levels and has been widely employed [8, 9]. Second, electrophilic compounds act as GSH‐depleting agents by covalently reacting with the thiol groups of GSH, thereby lowering its intracellular concentration [10, 11]. The third approach focuses on inhibiting GSH biosynthesis within the body, aiming to downregulate the production of this tripeptide through interference with its metabolic pathway [12, 13]. While these strategies have demonstrated efficacy in cancer therapy [14, 15], a critical unresolved question remains. This question concerns whether GSH, serving as a key homochiral biothiol and essential regulator in mammalian tissues, undergoes enantioselective interactions with chiral nanomaterials and how such enantioselectivity could impact the efficacy and specificity of cancer therapies.

Chirality, an intrinsic property in nature and biology, plays an essential role in numerous physiological processes, underpinning the exquisite stereoselectivity seen in enzyme specificity, cell signaling, and immune responses [16, 17, 18, 19, 20, 21, 22]. Understanding the enantioselective interactions between chiral nanomaterials and homochiral biomolecules is crucial for elucidating chiral biorecognition mechanisms and for developing targeted therapies and diagnostic tools in cancer treatment [23, 24]. The enantioselective interactions between chiral nanomaterials and biological systems, including biomolecules and cells, have attracted significant interest in biomaterial sciences and nanomedicine [25, 26, 27, 28]. Bioeffects of chirality have been demonstrated in protein adsorption [29], cell adhesion [30], proliferation and differentiation [31], cell phagocytosis [32, 33], and cell apoptosis [34]. Most reported chiral bioeffects stem from stereospecific interactions between chiral nanostructures and cell surface proteins [35, 36]. However, understanding how enantioselective interactions function in cancer therapies remains challenging due to the high complexity of the tumor microenvironment (TME), which poses significant hurdles for the design of chiral nanomaterials [37]. Metal nanoclusters (NCs), a critical class of nanomaterials, emerge as excellent candidates for chiral design in cancer therapy due to their atomically precise structures, exceptional structural tunability, superior assembly properties, and outstanding chiroptical characteristics [38, 39, 40, 41, 42, 43, 44, 45, 46]. To develop ideal chiral NCs with enantioselective effects for cancer therapy, three prerequisites are essential, (1) strong chiroptical activity and excellent biocompatibility to enable efficient cellular internalization, (2) robust enantioselective interactions with target biomarkers in cancer cells, and (3) favorable therapeutic efficacy derived from the enantioselective interactions.

In this study, we designed and fabricated a pair of silver nanoclusters (Ag₆NCs) with bilevel chirality as potent cancer therapeutic agents. This pair of nanoclusters enables enantioselective interactions with GSH in tumor cells, significantly enhancing therapeutic efficacy (Scheme 1). The chiral Ag₆NCs were further modified via PEGylation to improve biocompatibility, resulting in D‐Ag₆SP₆@PEG or L‐Ag₆SP₆@PEG nanoparticles [47]. Notably, these nanoparticles can selectively deplete intracellular GSH in tumor cells through ligand‐exchange reactions, subsequently forming Ag(I)‐GSH coordination supramolecular polymers that spontaneously assemble into microfibers in situ. Compared to its L‐type enantiomer, D‐Ag₆SP₆@PEG exhibits stronger GSH‐binding affinity, leading to enhanced intracellular GSH depletion. This disparity induced more severe mitochondrial mechanical damage through microfiber formation, consequently elevating ROS levels and significantly activating apoptosis and ferroptosis pathways in tumor cells. Furthermore, in vitro and in vivo studies have demonstrated that D‐Ag₆SP₆@PEG nanoparticles exhibit enhanced tumor‐killing efficacy compared with L‐Ag₆SP₆@PEG. Overall, our findings demonstrate the potential of this chirality‐dependent GSH depletion strategy for the development of precision cancer therapy, and also expand the applications of chiral nanomaterials in biomedicine.

SCHEME 1.

Illustrative schematics of enantioselective cancer therapy using Ag₆NCs with bilevel chirality. (a) Enantioselective GSH depletion via ligand exchange with chiral Ag₆NCs. (b) Mechanism diagram of D‐Ag₆SP₆@PEG inducing apoptosis and ferroptosis in tumor cells by depleting intracellular GSH and damaging mitochondria.

2. Results and Discussion

2.1. Synthesis and Characterization of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG

D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles were rationally designed and carefully prepared according to the procedure illustrated in Figure 1a. First, the ligands of D‐SP and L‐SP were synthesized according to the reported literature [48, 49]. The reaction of either D‐SP or L‐SP with CH₃COOAg was carried out in tetrahydrofuran (THF) at room temperature to obtain the D‐Ag₆SP₆ and L‐Ag₆SP₆ nanoclusters, respectively (details are provide in the Supporting Information). The chemical composition of D‐Ag₆SP₆ and L‐Ag₆SP₆ nanoclusters was characterized using high‐resolution electrospray ionization mass spectrometry (ESI‐HRMS). As shown in Figure 1b, the signal located at m/z = 1617.6788 corresponds to [D‐Ag₆SP₅]⁺. The corresponding signal for its enantiomer, [L‐Ag₆SP₅]⁺, is presented in Figure S1. Both as‐synthesized D‐Ag₆SP₆ and L‐Ag₆SP₆ exhibited good dispersity in transmission electron microscopy (TEM) images, with an average diameter of approximately 2 nm when suspended in ethanol (Figure S2). To gain insight into their precise chemical structures, single crystal x‐ray diffraction (SCXRD) was performed. As depicted in Figure S3, the distortion of the Ag₆ kernel was induced by the capping chiral ligands of D‐SP or L‐SP. The six phenyl groups on the ligands are arranged in three staggered pairs, adopting a propeller‐like conformation that imparts hierarchical chirality to the resulting Ag₆ nanoclusters at both the molecular and nanoscale levels.

FIGURE 1.

Synthesis and characterization of D/L‐Ag₆SP₆@PEG. (a) Schematic illustration for the synthesis of D/L‐Ag₆SP₆@PEG. (b) Positive ion mode ESI‐MS of D‐Ag₆SP₆ dissolved in DMF. The cyan line represents the measured isotopic distribution pattern, while the pink line shows the simulated isotopic distribution for the [D‐Ag₆SP₅]⁺ species. (c) SEM image of D‐Ag₆SP₆@PEG. (d) Hydrodynamic sizes of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG as measured by DLS. (e) CD, (f) PL, and (g) CPL spectra of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG in PBS (concentrations: 7.8 µM, pH = 7.4).

To improve the biocompatibility of D‐Ag₆SP₆ and L‐Ag₆SP₆, water‐soluble nanoparticles D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG were developed through a PEGylation strategy using DSPE‐PEG₂₀₀₀. Scanning electron microscope (SEM) images illustrated that the resultant D‐Ag₆SP₆@PEG nanoparticles had a uniform spherical morphology (Figure 1c). Following PEG modification, the surface potential of D‐Ag₆SP₆ was observed to decrease from 4.7 to −44.7 mV, which further confirmed the successful preparation of D‐Ag₆SP₆@PEG nanoparticles (Figure S4a). The zeta potential of L‐Ag₆SP₆@PEG is −43.6 mV, which is close to that of D‐Ag₆SP₆@PEG (Figure S4b). Dynamic light scattering (DLS) indicated that the average hydrodynamic sizes of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles were 85.5 and 84.8 nm, respectively (Figure 1d). Additionally, DLS measurements further revealed that D‐Ag₆SP₆@PEG showed no significant change in its average hydrodynamic size under different physiological environments, including phosphate‐buffered saline (PBS) and DMEM medium, indicating good stability (Figure S5).

The chirality of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles was confirmed by circular dichroism (CD) spectroscopy. As shown in Figure S6, the CD spectrum of D‐Ag₆SP₆ exhibited a negative Cotton effect at 270 nm, while L‐Ag₆SP₆ showed a mirror‐image CD curve corresponding to that of D‐Ag₆SP₆. After PEGylation, the spectra of D‐Ag₆SP₆@PEG displayed an enhanced negative CD signal at 275 nm, and L‐Ag₆SP₆@PEG exhibited enhanced positive CD absorption at 275 nm. The results unequivocally showed that modifying both the D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoclusters with DSPE‐PEG₂₀₀₀ induced ordered aggregation, thereby enhancing their chiroptical performance (Figure 1e). Additionally, both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG exhibited intense orange emission around 590 nm in PBS due to the aggregation‐induced emission (AIE) properties of the D‐Ag₆SP₆ and L‐Ag₆SP₆ nanoclusters (Figure 1f) [50]. Their CPL characteristics were measured in Figure 1g, where both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG presented remarkable CPL response around 590 nm corresponding to their photoluminescence (PL) shown in Figure 1f. The results clearly indicated that the two nanoparticles showed strong chiroptical activity and outstanding PL performance, making them suitable for subsequent biological research.

2.2. Enantioselective GSH Depletion of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG

Given the favorable chiroptical properties of Ag₆NCs and the thiophilicity of silver(I) ions [51], the depletion of GSH by D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG was systematically evaluated. As illustrated in Figure S7, the CD intensity at 275 nm for D‐Ag₆SP₆@PEG increased from −24.00 to −0.34 mdeg with the increasing in GSH concentration, thereby confirming the depletion of GSH. The differences in the CD response of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG to GSH were subsequently investigated. As depicted in Figure 2a, the CD intensity at 275 nm exhibited a significant increase, while at 360 nm, a trend of initial weakening followed by strengthening was observed in the D‐Ag₆SP₆@PEG and GSH systems. Moreover, the UV–visible (vis) absorption peaks at both 275 nm and 360 nm were enhanced within 10 min. The reactions involving L‐Ag₆SP₆@PEG and GSH are detailed in Figure S8. These findings indicate the rapid responsiveness and depletion capability of both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG toward GSH. Notably, D‐Ag₆SP₆@PEG demonstrated a faster reaction rate with L‐GSH (K ₁ = 0.09399), which was 1.45 times higher than that of L‐Ag₆SP₆@PEG (K ₂ = 0.06459) (Figure 2b). Based on these observations, the association rate constants (K _on) were determined for the reactions of D‐Ag₆SP₆@PEG with GSH (K _on = 7.23 × 10⁵ M⁻¹S⁻¹) and L‐Ag₆SP₆@PEG with GSH (K _on = 4.97 × 10⁵ M⁻¹S⁻¹), indicating faster binding of D‐Ag₆SP₆@PEG to GSH than that of L‐Ag₆SP₆@PEG. To further verify the different capabilities of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG to deplete GSH, the binding constants (K _a) of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG with GSH were calculated using the fluorescence titration method (Figures 2c and S9). The K _a value for L‐Ag₆SP₆@PEG with L‐GSH was determined to be 2.47 × 10⁷ M⁻¹, whereas D‐Ag₆SP₆@PEG with L‐GSH exhibited a significantly higher K _a value of 1.33 × 10⁸ M⁻¹. This result indicates that D‐Ag₆SP₆@PEG has a stronger binding affinity to L‐GSH than L‐Ag₆SP₆@PEG. The distinct binding affinities can be attributed to the better spatial matching of chirality between D‐Ag₆SP₆@PEG and L‐GSH, as well as the enhanced structural stability of the D‐Ag₆SP₆ + L‐GSH complex.

FIGURE 2.

Enantioselective GSH depletion of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. (a) CD spectra (top) and corresponding UV–vis spectra (bottom) of D‐Ag₆SP₆@PEG (7.8 µM) following treatment with GSH (80 µM) over various time intervals in PBS (pH = 7.4). (b) Linear correlation between the reaction time of D‐Ag₆SP₆@PEG (cyan) and L‐Ag₆SP₆@PEG (rose‐red) with GSH and the change in absorbance intensity (ΔAbs = Abs−Abs₀, where λ _dt = 275 nm). (c) Double‐logarithmic regression plots depicting the quenching effect of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG (7.8 µM) exposed to varying concentrations of GSH (0−100 µM). (d) Corresponding constants derived from the data in (b) and (c). K _a, K _on, and K _off denote the binding constants, association rate constants, and dissociation rate constants between D‐Ag₆SP₆@PEG/L‐Ag₆SP₆@PEG and GSH, respectively. (e) Illustration describing the mechanism responsible for the differential GSH depletion when interacting with D‐Ag₆SP₆@PEG or L‐Ag₆SP₆@PEG. 2D pseudo‐color images of (f) D‐Ag₆SP₆:GSH at a molar ratio of 1:1 and (g) L‐Ag₆SP₆:GSH at a molar ratio of 1:1. Early stage of absorption spectra of (h) D‐Ag₆SP₆:GSH at a molar ratio of 1:1, excited at 355 nm with a fluence of 0.2 µJ, and (i) L‐Ag₆SP₆:GSH at a molar ratio of 1:1, an excited at 355 nm with a fluence of 0.2 µJ. (j) Energy‐level diagram illustrating the photo‐physics of D/L‐Ag₆SP₆ and their complexes with GSH.

To verify the hypothesis, the dissociation rate constants (K _off) for the reactions involving D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG with GSH were determined using K _on and K _a. As depicted in Figure 2d, the K _off value for D‐Ag₆SP₆@PEG with GSH was 5.44 × 10⁻³ s⁻¹, while that for L‐Ag₆SP₆@PEG with GSH was 2.01 × 10⁻² s⁻¹. These values confirm that the reaction between D‐Ag₆SP₆@PEG and GSH exhibits a faster association rate and a slower dissociation rate compared to that of L‐Ag₆SP₆@PEG and GSH, resulting in enantioselective depletion of GSH (Figure 2e). Furthermore, femtosecond and nanosecond transient absorption (fs‐ and ns‐TA) spectra were recorded for the reactions of GSH with both D‐Ag₆SP₆ and L‐Ag₆SP₆. As depicted in Figure 2f,g, three distinct excited states were observed for both D‐Ag₆SP₆ + GSH and L‐Ag₆SP₆ + GSH. These sates were attributed to the excited state S₁ of Ag NCs at 495 nm, the excited states S’₁ of D‐Ag₆SP₆ + GSH at 614 nm, and the excited state S″₁ of L‐Ag₆SP₆ + GSH at 688 nm, as identified through excited‐state absorption experiments using 355 nm pump excitation (Figure 2h,i). Based on fs‐TA spectra, the energy scheme governing the photophysics of D‐Ag₆SP₆ and L‐Ag₆SP₆ as well as their complexes with GSH was elucidated. Both D‐Ag₆SP₆ and L‐Ag₆SP₆ were initially excited from the ground state (S₀) to higher excited state (S_n), then relaxed to the lowest excited singlets (S₁) through thermalization, and finally returned to the ground state (S₀) via internal conversion (Figure S10). The energy gap (S₀−S₁) of D‐Ag₆SP₆ + GSH (2.02 eV) was found to be larger than that of L‐Ag₆SP₆ + GSH (1.80 eV), suggesting a more stable complex formed by D‐Ag₆SP₆ + GSH. This finding serves as indirect evidence supporting the enantioselective depletion of GSH by D‐Ag₆SP₆ and L‐Ag₆SP₆ (Figure 2j).

2.3. GSH‐Depletion‐Mediated Self‐Assembly of D‐Ag₆SP₆@PEG

To investigate the morphological changes of D‐Ag₆SP₆@PEG at different time points after GSH treatment, TEM and SEM were employed for characterization and analysis. Upon mixing with 80 µM GSH, the D‐Ag₆SP₆@PEG nanoparticles gradually formed micrometer‐scale nanoribbons over 6 min, and the entire assembly process was completed in 15 min. With further incubation up to 24 h, the aggregates grew and ultimately formed microscale fibers (Figures 3a and S11). To elucidate the process of GSH‐depletion‐mediated self‐assembly by D‐Ag₆SP₆@PEG, DLS measurements were conducted at various time intervals. As shown in Figure 3b, the hydrodynamic diameters of the nanoparticles increased from an initial 83 nm to sizes ranging from 249 up to 1275 nm within 15 min, indicating their aggregation into larger nanostructures. Conversely, when D‐Ag₆SP₆@PEG was incubated with PBS at pH 7.4 for 24 h (Figure S12), the DLS data revealed minimal changes, underscoring the high stability of D‐Ag₆SP₆@PEG under standard physiological conditions. The GSH‐induced transition was further confirmed by PL measurements. As shown in Figure S13, the emission intensity at 580 nm decreased significantly within 6 min, demonstrating that the D‐Ag₆SP₆@PEG nanoparticles exhibit a rapid and pronounced response to GSH.

FIGURE 3.

GSH‐driven assembly of D‐Ag₆SP₆@PEG. (a) TEM images of D‐Ag₆SP₆@PEG after treatment with GSH (80 µM) over different time periods in PBS (pH = 7.4). (b) Hydrodynamic sizes of D‐Ag₆SP₆@PEG after treatment with GSH for 0, 5, and 15 min. (c) Schematic illustration of the assembly processes of D‐Ag₆SP₆@PEG in the presence of GSH. (d) CD spectra (top) and corresponding UV–vis spectra (bottom) of D‐Ag₆SP₆@PEG after treatment with GSH for 10 min in PBS buffer at pH = 5. (e) pH‐responsive changes in ΔAbs (λ _dt = 275 nm) during the ligand‐exchange between D‐Ag₆SP₆@PEG and GSH. Data are represented as mean ± s.e.m. (standard error of the mean, n = 3). (f) Comparative analysis of GSH depletion capabilities of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG (9 µM) were incubated with GSH (1 mM) at room temperature for different time intervals (2, 4, 6, 8, 10, and 12 min), respectively. Data are represented as mean ± s.e.m. (n = 3).

To confirm the existence of the Ag‐GSH complex, we have performed additional high‐resolution mass spectrometry (MS) experiments to monitor the changes in the MS signals of D‐Ag₆SP₆ clusters incubated with different concentrations of GSH (0, 15, 30, and 50 µM) (Figure S14a). After dissolving D‐Ag₆SP₆ clusters in DMF solution without GSH incubation (the cyan curve in Figure S14a), the MS spectrum of D‐Ag₆SP₆ clusters exhibited a predominant peak at m/z 1617.4517, which corresponds to the isotopic distribution pattern of [D‐Ag₆SP₅]⁺ (Figure S14b). Upon adding 15 µM GSH to the D‐Ag₆SP₆ cluster solution and allowing full incubation for 10 min (see the pink curve in Figure S14a), two new peaks emerged at m/z 2448.7655 and 1216.9465, corresponding to the isotopic distribution patterns of [D‐Ag₆SP₃(GSH)₄]⁺ and [D‐Ag₃SP₃GSH]⁺, respectively (Figure S14c). When 30 µM GSH was added and the mixture was incubated for 10 min (see the light pink curve in Figure S14a), the [D‐Ag₆SP₃(GSH)₄]⁺ signal disappeared entirely, replaced by two prominent peaks at m/z 1216.9465 and 829.9375, which correspond to the isotopic distributions of [D‐Ag₃SP₃GSH]⁺ (Figure S14d) and [Ag₂(GSH)₂]⁺ (Figure S14e), respectively. Increasing the GSH concentration to 50 µM (see the blue curve in Figure S14a) and allowing thorough incubation resulted in the disappearance of the [D‐Ag₃SP₃GSH]⁺ signal, with mainly the [Ag₂(GSH)₂]⁺ signal remaining detectable. This indicates that D‐Ag₆SP₆ clusters were fully dissociated into D‐SP molecules and Ag₂(GSH)₂ complexes after incubation with sufficient GSH. Based on the aforementioned findings, Figure 3c presents a schematic illustration of the self‐assembly mechanism of D‐Ag₆SP₆@PEG upon GSH triggering. Upon the addition of GSH, D‐Ag₆SP₆@PEG nanoparticles dissociate and first form Ag(I)‐GSH complexes through a ligand‐exchange reaction. This enantioselective recognition and interaction between D‐Ag₆SP₆ and GSH was further confirmed by ¹H NMR titration experiments (Figure S15). These complexes then assemble into Ag(I)‐GSH‐based coordination supramolecular polymers (CSPs), facilitated by intermolecular hydrogen bonding interactions [52]. After adequate aging, the resulting CSPs aggregate into microscale fibers.

To investigate the impact of the weakly acidic tumor microenvironment, we studied the GSH‐induced assembly of D‐Ag₆SP₆@PEG under different pH conditions (5.0, 6.5, and 7.4) by monitoring the time‐dependent changes in its UV absorption at 275 nm (Figure S16). As clearly illustrated in Figure 3d,e, the fastest GSH‐response rate and the highest concentration of Ag(I)‐GSH CSPs were detected under pH 5.0 conditions, suggesting that mildly acidic environments can accelerate the GSH‐induced structural transition of D‐Ag₆SP₆@PEG into Ag(I)‐GSH CSPs. As a control, the UV–vis absorption and CD spectra of D‐Ag₆SP₆@PEG without the addition of GSH in PBS at different pH levels were measured (Figure S17). When the pH value increased from 3.0 to 11.0, the CD spectra and UV–vis absorption spectra of D‐Ag₆SP₆@PEG exhibited no significant changes, indicating that solely adjusting the pH value had a negligible effect on the stability on the D‐Ag₆SP₆@PEG. Subsequently, we investigated the GSH‐depleting capabilities of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. D‐Ag₆SP₆@PEG exhibited a higher level of GSH consumption compared to L‐Ag₆SP₆@PEG (Figure 3f). These results demonstrate that GSH depletion triggers the rapid self‐assembly of D‐Ag₆SP₆@PEG nanoparticles into micrometer‐scale fibers through the formation of CSPs, a process that is accelerated under weakly acidic conditions typical of the tumor microenvironment.

2.4. Cell Uptake and Cytotoxicity of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG

To investigate the optimal time for cellular uptake of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG, we employed CT26 cells, a commonly used mouse colon cancer cell model. The uptake processes of these nanoparticles were monitored via PL spectroscopy, which capitalizes on the strong PL properties of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. As incubation time extended, the PL signal of D‐Ag₆SP₆@PEG gradually diminished, confirming their successful cellular uptake (Figures 4a and S18). As a control, the PL intensity remained nearly constant in both the CT26/DMEM and DMEM/D‐Ag₆SP₆@PEG systems. Moreover, the PL intensity profile of L‐Ag₆SP₆@PEG closely matched that of D‐Ag₆SP₆@PEG and plateaued within 8 h, indicating similar cellular uptake kinetics for both nanoparticles (Figure S19). It was further evidenced by quantifying the intracellular silver (Ag) levels using inductively coupled plasma mass spectrometry (ICP‐MS) (Figure 4b). The intracellular Ag content reached a maximum at 10 h and did not increase thereafter. Collectively, the results indicate that both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG were efficiently internalized by CT26 cells, with no difference in their cellular uptake behavior.

FIGURE 4.

In vitro assessment of cancer therapeutic effects of D‐Ag₆SP₆@PEG nanoparticles. (a) Quantitative analysis of PL intensity (λ _dt = 580 nm) from different culture systems. Data are presented as mean ± s.e.m. (n = 4). (b) Quantification of silver uptake in CT26 cells evaluated by ICP‐MS (n = 3). (c) Cell viability of CT26 cells incubated with D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG for 24 h. Data are presented as mean ± s.e.m. (n = 4). (d) Representative fluorescence images of Calcein‐AM (live cells, green) and PI (dead cells, red) co‐stained CT26 cells following 24 h exposure to PBS, L‐SP@PEG, D‐SP@PEG, L‐Ag₆SP₆@PEG, or D‐Ag₆SP₆@PEG (all at 7.2 µM, except PBS). (e) Flow cytometric analysis of CT26 cells apoptosis following different treatments. (f) Photographs showing the colony formation of CT26 cells subjected to various treatments. (g) Statistical analysis of colony formation efficiency in CT26 cells following different treatments. Data are represented as mean ± s.e.m. (n = 3). Statistical significance was determined using one‐way ANOVA followed by Tukey's post‐hoc test. Significance levels are indicated as follows: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

The cytotoxicity of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG was evaluated in both cancer cells and normal cells through a standard CCK‐8 assay, following a 24 h exposure to various concentrations of the nanoclusters or their ligand controls (D‐SP@PEG and L‐SP@PEG). To verify the difference in intracellular GSH levels between the two cell lines, we quantified GSH concentrations in IEC‐6 and CT26 cells using a GSH assay kit. As shown in Figure S20, the GSH level in CT26 cells was approximately threefold higher than that in IEC‐6 cells. This marked difference supports the selective responsiveness of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles. For IEC‐6 cells, neither D/L‐SP@PEG nor D/L‐Ag₆SP₆@PEG exhibited obvious cytotoxicity (Figure S21). In contrast, as shown in Figure 4c, CT26 cell viability gradually decreased with increasing concentrations of D‐Ag₆SP₆@PEG or L‐Ag₆SP₆@PEG. The half‐maximal inhibitory concentration (IC₅₀) was calculated to quantify the cytotoxicity of L‐Ag₆SP₆@PEG, which was determined to be 5.04 ± 0.17 µM for CT26 cells. In comparison, D‐Ag₆SP₆@PEG exhibited superior inhibitory activity against CT26 cells, with an IC₅₀ of 4.05 ± 0.03 µM. Significant differences in cell viability between the two enantiomers were observed at concentrations above 5.4 µM, and this disparity became more pronounced at higher concentrations. These findings suggest that D‐Ag₆SP₆@PEG exerts greater inhibitory efficacy against cancer cells than L‐Ag₆SP₆@PEG.

We further investigated the cytotoxicity of both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG in other cell lines, including non‐small cell lung cancer cells (A549), human cervical carcinoma cells (HeLa), and normal human bronchial epithelial cells (BEAS‐2B) (Figures S22 and S23). The results showed that the IC₅₀ values for L‐Ag₆SP₆@PEG were 6.17 ± 0.14 µM for A549 cells and 8.29 ± 0.09 µM for HeLa cells, whereas D‐Ag₆SP₆@PEG exhibited enhanced inhibitory efficacy with IC₅₀ values of 5.10 ± 0.09 µM for A549 cells and 5.58 ± 0.14 µM for HeLa cells. Conversely, over 98% of BEAS‐2B cells remained viable across all tested concentrations. Collectively, these results indicate that both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG exert potent antitumor activity while displaying minimal cytotoxicity toward normal cells. Notably, D‐Ag₆SP₆@PEG demonstrated superior inhibitory activity against cancer cells compared with its L‐enantiomer, highlighting the critical role of cluster chirality in cancer therapy.

The cancer therapeutic effects of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG were further evaluated by live/dead cell staining analysis using Calcein‐AM and propidium iodide (PI), as well as by flow cytometry using Annexin V‐FITC and PI staining in vitro. As shown in Figure 4d, after 24 h of incubation, red fluorescence indicative of cell death was observed in both the D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG treatment groups. However, the fluorescence signal of the D‐Ag₆SP₆@PEG group was significantly stronger than that of the L‐Ag₆SP₆@PEG group, indicating superior killing efficacy toward CT26 cells. The live/dead staining images and corresponding red fluorescence intensity analysis of A549 cells under different treatments further confirmed that D‐Ag₆SP₆@PEG exerted a more potent cell‐killing effect (Figure S24). Meanwhile, in HeLa cells, the red signal intensity gradually increased with elevated concentrations of D‐Ag₆SP₆@PEG, indicating enhanced cytotoxicity (Figure S25). Flow cytometry analysis indicated that the apoptosis rate of CT26 cells in the D‐Ag₆SP₆@PEG group was 1.74‐fold higher than in the L‐Ag₆SP₆@PEG group (Figures 4e and S26). Similarly, in A549 cells, D‐Ag₆SP₆@PEG also induced the highest apoptosis rate compared to L‐Ag₆SP₆@PEG and control groups (Figure S27). A colony formation assay was further performed to assess the therapeutic effects of the two nanoclusters. As shown in Figure 4f,g, D‐Ag₆SP₆@PEG suppressed colony formation by approximately 86% in CT26 cells, whereas L‐Ag₆SP₆@PEG inhibited it by approximately 66.8%. Similarly, consistent results were observed in A549 cells, where the colony formation rate in the D‐Ag₆SP₆@PEG group was significantly lower than that in the L‐Ag₆SP₆@PEG group (Figure S28). These results confirm that both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG possess significant anti‐cancer activity, with D‐Ag₆SP₆@PEG showing superior efficacy compared to L‐Ag₆SP₆@PEG.

2.5. Mechanism of Cancer Cell Death Induced by D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG

We sought to investigate the cellular mechanisms underlying the potent anti‐tumor effects of D‐Ag₆SP₆@PEG. Based on previous findings, both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG nanoparticles can assemble into Ag(I)‐GSH‐based CSPs upon encountering intracellular GSH. We propose that this supramolecular assembly directly induces mechanical damage to mitochondria, consequently increasing intracellular ROS levels. Meanwhile, the assembly process depletes intracellular GSH, which in turn leads to the inactivation of glutathione peroxidase 4 (GPX4). Elevated ROS combined with GPX4 inactivation ultimately cooperate to trigger apoptosis and ferroptosis in tumor cells (Figure 5a). To assess intracellular ROS generation, 2ʹ,7ʹ‐Dichlorodihydrofluorescein diacetate (DCFH‐DA) served as the detection probe. As shown in Figure 5b, a notable increase in fluorescence was observed in the FITC channel for cells treated with D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. Statistical analysis revealed that the mean fluorescence intensity in the D‐Ag₆SP₆@PEG group was significantly higher than in the other groups, approximately 1.3 times that of L‐Ag₆SP₆@PEG group and 2.6 times that of the control group (Figure 5c). Fluorescence microscopy images of DCFH‐DA‐stained CT26 cells subjected to different treatments showed strong green fluorescence in the D‐Ag₆SP₆@PEG group, followed by the L‐Ag₆SP₆@PEG group, whereas minimal fluorescence was observed in the PBS, D‐SP@PEG, and L‐SP@PEG groups (Figure 5d). Further evaluation in A549 and HeLa cell lines revealed that D‐Ag₆SP₆@PEG produced the highest ROS levels among all groups (Figures S29 and S30). These results confirm that D‐Ag₆SP₆@PEG induce significantly higher ROS production than L‐Ag₆SP₆@PEG across the tested cell lines.

FIGURE 5.

Mechanism of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG‐induced cell death. (a) Schematic illustration of the enantioselective cancer therapy mediated by Ag₆NCs. (b) Flow cytometry analysis assessing the production of ROS in CT26 cells after different treatments. (c) Corresponding mean fluorescence intensity statistics from the flow cytometry analysis of ROS production in CT26 cells. Data are represented as mean ± s.e.m. (n = 3). (d) Fluorescence microscope images of CT26 cells stained with DCFH‐DA after treatment with PBS, L‐SP@PEG, D‐SP@PEG, L‐Ag₆SP₆@PEG, or D‐Ag₆SP₆@PEG (all at 7.2 µM, except PBS). (e) Images of CT26 cells stained with mBBr after different treatments. (f) Measurement of intracellular GSH concentration in CT26 cells after different treatments. Data are represented as mean ± s.e.m. (n = 3). (g) Western blotting analysis of cleaved Caspase‐3 expression in CT26 cells after different treatments. (h) Western blotting analysis of GPX4 expression in CT26 cells after different treatments. (i) Visualization of LPO in CT26 cells stained with C11‐BODIPY^581/591 after different treatments. (j) Representative TEM images of CT26 cells incubated with D‐Ag₆SP₆@PEG for different time points (yellow solid triangle: mitochondria; pink solid triangle: Ag(I)‐GSH aggregates; red dotted circle: D‐Ag₆SP₆@PEG). Statistical significance was determined using one‐way ANOVA followed by Tukey's post‐hoc test. Significance levels are indicated as follows: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

To verify that the inhibitory effects of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG on cancer cells are associated with GSH depletion, CT26 cells were stained with Thiol‐Tracker following various treatments. The D‐SP@PEG and L‐SP@PEG groups showed fluorescence intensity similar to that of the PBS group, indicating negligible effects on intracellular GSH levels (Figures 5e and S31). In contrast, cells treated with D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG exhibited significantly decreased fluorescence, demonstrating that both nanoclusters could efficiently deplete intracellular GSH. Notably, D‐Ag₆SP₆@PEG displayed stronger GSH‐depletion capacity in CT26 cells than L‐Ag₆SP₆@PEG. To further validate the qualitative observations, intracellular GSH levels in CT26 cells were quantitatively determined using a reduced GSH assay kit. As shown in Figure 5f, the concentration of intracellular GSH in untreated CT26 cells was measured as 5.39 ± 0.13 mM. Treatment with L‐Ag₆SP₆@PEG decreased intracellular GSH to 2.77 ± 0.16 mM, whereas D‐Ag₆SP₆@PEG induced a more pronounced reduction, with GSH levels dropping to 2.16 ± 0.08 mM. Thus, these results confirm that both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG can efficiently deplete GSH in cancer cells, with the D‐enantiomer demonstrating higher depletion efficiency than its L‐counterpart.

To better elucidate the detailed mechanism underlying tumor cell death, the ROS burst‐triggered apoptotic pathway was verified by Western blotting. As depicted in Figure 5g, compared with the PBS, L‐SP@PEG, and D‐SP@PEG groups, both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG treatments significantly upregulated the expression of cleaved caspase‐3, a canonical apoptosis‐related marker. Moreover, D‐Ag₆SP₆@PEG induced a more pronounced upregulation than L‐Ag₆SP₆@PEG. GSH depletion impairs the activity of GPX4, a key lipid repair enzyme, thereby promoting ferroptosis in tumor cells [53]. Figure 5h showed that treatment with D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG markedly downregulated GPX4 expression relative to the other three control groups, while D‐Ag₆SP₆@PEG exerted a stronger inhibitory effect on GPX4 than its L‐enantiomer. Furthermore, extensive GSH depletion and excessive ROS accumulation can lead to lipid peroxidation (LPO) in cancer cells [54]. We subsequently evaluated LPO levels in CT26 cells using C11‐BODIPY^581/591, a specific fluorescence probe whose emission spectrum shifts from 590 nm (red) to 510 nm (green) upon oxidation by LPO [55]. As shown in Figure 5i, both D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG groups exhibited enhanced green fluorescence intensity relative to the other groups, confirming LPO accumulation and ferroptosis induction mediated by these chiral AgNCs. Notably, D‐Ag₆SP₆@PEG displayed a significantly weaker red fluorescence signal and a stronger green fluorescence signal than L‐Ag₆SP₆@PEG, indicating its superior capacity to induce ferroptosis in tumor cells.

To provide direct evidence for the involvement of ferroptosis, we performed rescue experiments using ferrostatin‐1 (Fer‐1), a specific inhibitor of ferroptosis. As shown in Figure S32, CCK‐8 assay results demonstrated that pretreatment with Fer‐1 significantly attenuated cell death induced by D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG. Specifically, Fer‐1 treatment reduced the cell death rate from 76.75% to 30.50% in the D‐Ag₆SP₆@PEG group and from 56.25% to 15.75% in the L‐Ag₆SP₆@PEG group. Importantly, C11‐BODIPY^581/591 staining showed that co‐treatment with Fer‐1 completely abolished the LPO accumulation induced by both enantiomers, restoring the red fluorescence signal to levels comparable to those in the control group (Figure S33). Collectively, these findings validate the occurrence of ferroptosis during cell damage induced by the two chiral nanoparticles. The D‐enantiomer displays enhanced ferroptosis‐inducing capacity, and ferroptosis acts as a key pathway participating in their antitumor activity.

As previously verified, D‐Ag₆SP₆@PEG nanoparticles assemble into nanofibrous aggregates composed of Ag(I)‐GSH‐based CSPs in GSH‐containing solution, followed by the formation of microfibers. To investigate their intracellular self‐assembly and impacts on cell morphology and structure, biological transmission electron microscopy (bio‐TEM) was applied for morphology observation. At the initial untreated stage (0 h), intact mitochondrial cristae (indicated by yellow triangles) were clearly observed (Figures 5j and S34), indicating intact mitochondrial structure and normal physiological function. After 12 h of incuabtion, massive intracellular internalization of D‐Ag₆SP₆@PEG nanoparticles (indicated by red dotted lines) was observed. Meanwhile, the nanoparticles underwent GSH‐mediated intracellular ligand exchange to generate Ag(I)‐GSH complexes. These complexes subsequently self‐assembled into nascent fibrous CSPs (marked by pink triangles). Accompanied by this process, early mitochondrial structural abnormalities emerged, suggesting the initiation of mitochondrial injury. At 24 h post‐treatment, these fibrous CSPs further developed into microfibers. In addition, mitochondrial damage was dramatically aggravated, with extensive cristae loss and disrupted mitochondrial morphology. Such typical mitochondrial lesions are recognized as key morphological features during ferroptosis progression.

Subsequently, we further evaluated mitochondrial dysfunction induced by D/L‐Ag₆SP₆@PEG during their intracellular assembly in cancer cells. The fluorescent probe JC‐1 (5,5ʹ,6,6ʹ‐tetrachloro‐1,1ʹ,3,3ʹ‐tetraethylbenzimidazolocarbocyanine iodide) was employed to monitor mitochondrial membrane potential (MMP). JC‐1 forms red‐fluorescent aggregates in mitochondria with normal membrane potential, and dissociates into green‐fluorescent monomers upon mitochondrial depolarization. As shown in Figure S35, treatment with D/L‐Ag₆SP₆@PEG clearly reduced MMP, and this change was more obvious in the D‐Ag₆SP₆@PEG group. However, the other three groups showed no obvious changes in fluorescence intensity. These results confirm that the assemblies formed by the interaction between D/L‐Ag₆SP₆@PEG and intracellular GSH can damage mitochondria and cause subsequent MMP reduction. Taken together, we conclude that these nanoparticles exert anti‐tumor effects mainly by consuming intracellular GSH and further self‐assembling into microfibrous aggregates. These aggregates trigger mitochondrial structural damage and excessive ROS accumulation, thereby inducing both apoptosis and ferroptosis in tumor cells. Importantly, D‐Ag₆SP₆@PEG shows higher binding capacity toward GSH than L‐Ag₆SP₆@PEG. It can efficiently consume intracellular GSH and complete intracellular self‐assembly at a faster rate, thus exhibiting much stronger cytotoxicity against tumor cells.

2.6. In Vivo Antitumor Activity Assessment

The in vivo anti‐tumor efficacy of D‐Ag₆SP₆@PEG nanoparticles was evaluated using a CT26 subcutaneous tumor model. When the average tumor volume reached approximately 60 mm³, the mice were randomly divided into five groups: PBS group, L‐SP@PEG group, D‐SP@PEG group, L‐Ag₆SP₆@PEG group, and D‐Ag₆SP₆@PEG group. Each group received intra‐tumoral injections every two days at a dosage of 10 mg kg⁻¹, with a total of five treatments (Figure 6a). As shown in Figure 6b, D‐SP@PEG and L‐SP@PEG exerted only minimal inhibitory effects on tumor growth. Notably, D‐Ag₆SP₆@PEG was superior to its L‐enantiomer in suppressing tumor growth, and demonstrated the strongest inhibitory effect among all treatment groups. Tumors in this group exhibited the smallest visible size and the lowest average weight, which were consistent with the tumor volume monitoring results (Figure 6c,d).

FIGURE 6.

In vivo antitumor efficacy of D‐Ag₆SP₆@PEG and L‐Ag₆SP₆@PEG in CT26 tumor model. (a) Schematic illustration of the treatment protocol used for the CT26 tumor model. (b) Relative tumor volume growth curves of different treatment groups. Data are represented as mean ± s.e.m. (n = 5). (c) Photographs of excised tumors following different treatments. (d) Weights of the tumors corresponding to those shown in (c). Data are represented as mean ± s.e.m. (n = 5). (e) Representative images of DCFH‐DA, mBBr, and GPX4 staining in tumor sections after different treatments. (f) Statistical analysis of the mean FL intensity for DCFH‐DA, mBBr, and GPX4 staining, respectively. Data are represented as mean ± s.e.m. (n = 3). (g) H&E and TUNEL staining of tumor slices after different treatments. Statistical significance was determined using one‐way ANOVA followed by Tukey's post‐hoc test. Significance levels are indicated as follows: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Following different treatments, we measured the levels of ROS, GSH, and GPX4 in mouse tumor slices, with DCFH‐DA, mBBr, and anti‐GPX4 antibody used as the corresponding probes. As shown in Figure 6e,f, the D‐Ag₆SP₆@PEG group displayed the strongest DCFH‐DA fluorescence signal, indicating excessive ROS accumulation in tumor tissues. Additionally, D‐Ag₆SP₆@PEG led to the most significant GSH depletion, as evidenced by weak red fluorescence in tissue sections. Furthermore, this treatment markedly downregulated GPX4 expression. These in vivo results are consistent with in vitro findings, further validating that the regulation of ROS, GSH, and GPX4 serves as a key mechanism for the potent antitumor efficacy of D‐Ag₆SP₆@PEG.

To further evaluate the antitumor efficacy of D‐Ag₆SP₆@PEG nanoparticles, we performed hematoxylin and eosin (H&E) and TdT‐mediated dUTP nick‐end labeling (TUNEL) staining on tumor slices from mice subjected to different treatments. As shown in Figure 6g, H&E stained images of D‐Ag₆SP₆@PEG‐treated tumors demonstrated a marked reduction in viable tumor cell density and obvious structural disorganization, indicating potent inhibition of tumor cell proliferation. TUNEL staining revealed extensively positive signals in the D‐Ag₆SP₆@PEG treatment group, characterized by brown or dark brown staining, suggesting remarkably enhanced DNA fragmentation and apoptotic cell death. We also assessed the biosafety profile of D‐Ag₆SP₆@PEG. During the treatment period, all mouse groups showed no obvious changes in body weights, suggesting negligible adverse effects from the administered nanoparticles (Figure S36). After 14 days of treatment, major organs (heart, liver, spleen, kidney, lung, and intestine) in the D‐Ag₆SP₆@PEG‐treated group presented intact histological structures, comparable to those in the PBS group (Figure S37). In addition, no significant differences were observed in complete blood count and biochemical parameters between the D‐Ag₆SP₆@PEG group and the PBS group (Figure S38). Overall, these results demonstrate that D‐Ag₆SP₆@PEG exerts the most potent inhibitory effect on subcutaneous CT26 tumors across all treatment groups, with no obvious systemic toxicity observed.

3. Conclusion

In summary, we have rationally designed and synthesized a pair of bilevel chiral silver nanoclusters, D‐Ag₆SP₆ and L‐Ag₆SP₆, and their PEGylated derivatives, which exhibit enantioselective recognition and binding toward GSH. Owing to its higher binding affinity, D‐Ag₆SP₆@PEG enables more efficient intracellular GSH depletion via a chirality‐dependent ligand‐exchange process. Upon GSH depletion, it triggers a sequential assembly cascade that facilitates the formation of Ag(I)‐GSH complexes, followed by in situ self‐assembly into CSPs and the eventual generation of microscale fibrous structures. These aggregates induce mechanical disruption of mitochondria, leading to a marked increase in ROS production and the downregulation of GPX4. Such combined effects collectively promote apoptosis and ferroptosis in tumor cells. In vivo studies employing a CT26 subcutaneous tumor model confirm that D‐Ag₆SP₆@PEG exerts superior antitumor efficacy relative to its L‐enantiomer, without inducing obvious systemic toxicity. This work not only presents a new class of chiral nanotherapeutics but also highlights the critical role of hierarchical chirality in the design of bioactive nanoclusters, offering a promising strategy for enantioselective cancer therapy.

Author Contributions

Xuejuan Wang: investigation, writing – original draft, data curation, writing – review and editing, and methodology. Xirui Wu: methodology, investigation, writing – original draft, and writing – review and editing. Yue Zhao: investigation, data curation, and writing – review and editing. Xiangyang Zhang: investigation, methodology, and data curation. Jianfeng Zhao: methodology and investigation. Miaolong Li: investigation. Guangbao Yang: writing – review and editing, writing – original draft, supervision, project administration, and funding acquisition. Guofeng Liu: conceptualization, writing – review and editing, writing – original draft, project administration, supervision, and funding acquisition.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: anie72823‐sup‐0001‐SuppMat.pdf.

Data Availability Statement

The data that supports the findings of this study are available in the Supporting Information of this article.