Polysaccharide-silica nanocarriers for multimodal detection and anti-apoptotic therapy in hypertensive heart disease

2.1. Materials and instrumentation

All reagents and solvents were commercially available and used as received without further purification. Fourier-transform infrared (FT-IR) spectra were collected using a Thermo Scientific Nicolet iS10 FT-IR spectrometer. UV–visible absorption measurements were carried out on a Shimadzu UV-2600 spectrophotometer. Fluorescence emission spectra were obtained with a Horiba Fluorolog-3 spectrofluorometer. Powder X-ray diffraction (PXRD) patterns were recorded on a Rigaku Ultima IV diffractometer (Cu Kα radiation, λ = 1.5418 Å). Scanning electron microscopy (SEM) images were captured using a Hitachi S-4800 microscope. Electrochemical measurements (cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS)) were conducted on a CHI 660E electrochemical workstation.

2.2. Synthesis of compound 1 and CP1

Using 1,3-cyclohexanedione (0.140 g, 1.00 mmol), ethyl cyanoacetate (0.131 g, 1.00 mmol), and p-isopropylbenzaldehyde (0.147 mL, 1.00 mmol) as starting materials, sodium carbonate (0.106 g, 1.00 mmol) and 20 mL of ethanol were added, and the mixture was stirred under reflux for 4 h. Through a one-pot Knoevenagel condensation–Michael addition–cyclization sequence, the target Compound 1 was obtained. After completion, the solvent was evaporated, the crude product was extracted with ethyl acetate, washed with water, dried over sodium sulfate, and concentrated. Finally, recrystallization from hot ethanol at 70°C afforded yellow needles of Compound 1 in approximately 78% yield (about 0.225 g).

Dissolve a mixture of 0.15 mmol Dy(NO₃)₃ and 0.4 mmol 4-(1-hydroxy-1,4-dihydropyridin-4-yl)benzoic acid (H₂L) in a 12 mL ethanol-water mixed solvent (volume ratio 1:1), and then add HNO₃ to adjust the pH to 5. The resulting mixture was heated in a 25 mL stainless steel vessel lined with polytetrafluoroethylene at 125°C for 60 h. Subsequently, the reactor was cooled to ambient temperature to obtain colorless block-shaped crystals. The yield was 44% (based on H₂L). Elemental analysis for CP1: Anal. Calcd. for C₂₄H₁₆DyN₃O₉ (%): C, 44.15; H, 2.47; N, 6.44. Found: C, 44.12; H, 2.45; N, 44.07. The single crystal data has been shown in Table 1.

2.3. Synthesis of 1-DASA-ATPMS@CP1@Eth

200 mg of DASA precursor (sodium 2,4‐bis(acetoxy)‐3,5‐dimethylphenolate polymer, ∼0.50 mmol repeating units) and 150 mg of Compound 1 (4‐isopropyl-1,3‐dioxoisoindole‐2‐amine, 0.50 mmol) were dissolved in 20 mL MeOH under N₂; 70 µL Et₃N (0.50 mmol) was added, and the mixture was stirred for 12 h. After solvent removal, the crude was extracted with DCM/water and dried to give 1-DASA (260 mg, 85% yield). Next, 250 mg of 1-DASA (0.33 mmol) was suspended in 20 mL MeOH under N₂, and 150 µL ATPMS (0.75 mmol) plus 50 µL Et₃N (0.36 mmol) were added. After 12 h of stirring, the product was washed and dried to yield 1-DASA-ATPMS (≈ 280 mg, 90% yield). Finally, 200 mg of 1-DASA-ATPMS was dispersed in 10 mL MeOH, then 50 mg CP1 and 25 mg Eth were each dissolved in 5 mL phosphate-buffered saline (PBS) (pH 7.4) and added dropwise under stirring (first CP1 for 4 h, then Eth for 6 h). The resulting product was centrifuged, washed twice with PBS, and lyophilized to afford 1-DASA-ATPMS@CP1@Eth as a light yellow powder (≈ 245 mg, ∼75% overall yield).

2.4. CCK-8 assay

H9c2 rat embryonic cardiomyocytes (ATCC, USA) were cultured in DMEM medium containing 10% fetal bovine serum (FBS, Gibco, USA) in an incubator at 37°C, 5% CO₂. The experimental groups were as follows. Normal control group. Cells in the model group were stimulated with 1 μM angiotensin II (Ang II) for 24 h to induce apoptosis. Cells in the Eth-NPs group were supplemented with 40 μg mL^-1 nanoparticles loaded with ethoxysanguinarine (Eth-NPs) based on the model group. Cells in the free drug group (Eth) were added 40 μg mL^-1 of free ethoxyhaematoxylin based on the model group. Blank nanoparticles group (Blank) was added blank nanoparticles (without drug) based on the model group. The drug treatment was carried out for 24 h, and the cell activity of each group was detected using CCK-8 kit (Dojindo, Japan) according to the manufacturer’s instructions. The experiment was repeated independently three times (n=3).

2.5. qPCR analysis

RNA in cells was isolated using RNA extraction kit (Beyotime, China), total RNA was reverse transcribed using the PrimeScript RT Kit (TaKaRa, Japan). The qPCR was performed using SYBR Green Master Mix (TaKaRa, Japan). The primer sequences were as follows: Bax, Forward: CAGCAGAGCCAGATGAAGAA; Reverse: GCCATCTTCTGGAGTGGATG. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH), Forward: ACCACAGTCCATGCCATCAC; Reverse: TCCACCACCCTGTTGCTGTA. The experiment was repeated independently three times (n=3).

2.6. Fluorescence lifetime and quantum yield

Fluorescence lifetime was measured using time-correlated single-photon counting (TCSPC) with excitation at 330 nm. Quantum yield was calculated using quinine sulfate (Φ = 0.54) as reference under identical optical conditions. Nanoparticle dispersions were diluted to ensure absorbance <0.1 at the excitation wavelength to minimize inner-filter effects.

3.1. Structure description of compound 1

The single crystal XRD data of compound 1 show that compound 1 crystallizes with the monoclinic crystal system with the space group of Cc. As shown in Figure 1(a), the molecule structure consists of two six-membered rings (ring A: C4-C8 and O3, and ring B: C6-C7 and C9-C12) and one benzene ring C: C13-C18. The atom C4 of ring A connects an ethyl formate group (-C1H₅C2O₂) via the bond of C3-C4. The C5 atom of the ring A connects an amino group (-N1H₂). Ring A and ring B connect each other through the sharing of a C-C bond from C6 and C7. The atom C8 of ring A connects ring C via the C13 atom. The atom C12 of ring B connects the O4 atom via a double bond of C=O. The atom C19 of ring C connects two methyl group (-C21H₃ and -C20H₃). Furthermore, the neighboring molecules of compound 1 are further connected by intermolecular hydrogen bonds (O4-H1E…N1: 2.1040Å) (Figure 1b) and van der Waals forces, which give a three-dimensional dense packing structure (Figure 1c).

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Figure 1.

(a) The molecule structure for compound 1; (b) The H-bonds between the adjacent molecules; (c) The dense packing molecule structure of compound 1.

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3.2. Structure description of CP1

The single crystal XRD data analysis for CP1 reveals that CP1 shows a 3D porous structure based on the dinuclear secondary building units (SBUs) and crystallizes in the monoclinic crystal system with the C1 space group. Structure analysis reveals that the asymmetric unit of CP1 contains one Dy(III) ion, two L⁻ ligands, and one nitrate ion. As shown in Figure 2(a), the Dy(III) ion adopts an eight-coordination by eight oxygen atoms, in which four oxygen atoms come from four carboxylic groups of four independent L⁻ ligands, two oxygen atoms come from two organic ligands, and two oxygen atoms from one nitrate ion (Figure 2b and 2c). In CP1, the L^- connects three Dy³⁺ ions through one carboxylate group and one oxygen atom. The carboxylic groups adopt the bridging coordination model (η²μ²χ²) to connect two Dy(III) ions and the oxygen atom of the L^- connect one Dy(III) ion (Figure 2b). The two eight-connected Dy(III) ions were connected by four carboxylic groups, four oxygen atoms, and two nitrate ions to produce a dinuclear SBUs (Figure 2d). The SBUs were further extended by the organic ligands to give a 3D porous structure (Figure 2e).

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Figure 2.

(a) The coordinated environment of Dy(III) ion in CP1 (Symmetry codes: #1: -x, y, 3/2-z; #2: -1/2+x, 1/2-y, 1/2+z; #3: 1/2-x, -1/2+y, 3/2-z); (b) The ligand coordination pattern for CP1; (c) Coordination geometries of Dy(III) ion; (d) The dinuclear SBU in CP1; (e) The 3D porous structure of CP1.

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3.3. Characterization of 1-DASA-ATPMS@CP1@Eth

Figure 3 presents the X‐ray diffraction (XRD) patterns and Fourier‐transform infrared (FTIR) spectra of 1‐DASA‐ATPMS@CP1 (blue) and its Eth loaded counterpart 1‐DASA‐ATPMS@CP1@Eth (orange), confirming successful Eth incorporation. In the XRD patterns (Figure 3a), both samples exhibit broad reflections attributable to the semi‐crystalline ATPMS framework and ordered arrangement of DASA ligands and CP1 units (notably near 2θ ≈ 8–10°, 15–17°, and 29–31°). Upon Eth loading, these characteristic peaks are retained but display reduced intensity and slight broadening, most pronounced around 2θ ≈ 9° and 31°, indicating that Eth molecules occupy the pore spaces or adhere to the surface, partially disrupting long‐range ordering without collapsing the mesostructure. Complementarily, FTIR spectra of 1‐DASA‐ATPMS@CP1 feature diagnostic bands at 2920–2850 cm⁻¹ (alkyl C–H stretches), ∼1720 cm⁻¹ (DASA ester C=O stretch), and 1510–1460 cm⁻¹ (aromatic C=C and triazole stretches), alongside Si–O–Si vibrations at 1080–1040 cm⁻¹ (Figure 3b). After Eth incorporation, new broad absorptions appear at 3380–3300 cm⁻¹ (O–H/N–H stretches) and 1600–1580 cm⁻¹ (aromatic C=C/N–H bends from Eth), the C=O stretch shifts slightly from 1720 to 1705 cm⁻¹ (consistent with hydrogen bonding between Eth and the DASA carbonyl), and subtle changes arise in the Si–O–Si region, reflecting minor framework distortion. In Figure 3c, the SEM image of an individual 1-DASA-ATPMS@CP1@Eth nanoparticle reveals a quasi-spherical architecture with a textured surface composed of densely arranged sub-400 nm protrusions. The particle exhibits an anisotropic size profile, measuring approximately 632 nm in the vertical direction and 785 nm along the horizontal axis, reflecting its slightly elliptical morphology and hierarchical surface structure. Together, these XRD, FTIR, and SEM changes unequivocally demonstrate that Eth has been successfully loaded into 1‐DASA‐ATPMS@CP1, yielding the final 1‐DASA‐ATPMS@CP1@Eth composite. In addition, the thermal stabilities of compound 1 and CP1 were further evaluated by thermogravimetric analysis (TGA), as shown in Figure 3d. Compound 1 exhibits an initial weight loss below 150°C, corresponding to the removal of physically adsorbed water and residual solvents, followed by a major decomposition stage between 250–450°C, reflecting the breakdown of its organic backbone. In contrast, CP1 shows a slightly earlier onset of thermal degradation, beginning around 200°C, and undergoes a broader and more gradual weight-loss process up to 600°C due to the combination of coordinated ligands and the thermally robust Dy–O framework. The remaining mass at 800°C is higher for CP1, consistent with the presence of thermally stable inorganic Dy-containing residues. These TGA results confirm that both compound 1 and CP1 possess sufficient thermal stability to withstand subsequent nanocomposite fabrication steps.

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Figure 3.

Characterization of 1-DASA-ATPMS@CP1@Eth: (a) PXRD, (b) FTIR, (c) SEM, and (d) TGA.

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3.4. The peroxidase-like activity of 1-DASA-ATPMS@CP1@Eth

Oxidative stress plays a central role in the pathogenesis of HHD: sustained elevated blood pressure leads to excessive accumulation of H₂O₂ within cardiomyocytes, which in turn triggers lipid peroxidation, protein oxidation, and DNA damage, ultimately causing cardiomyocyte apoptosis, fibrosis, and cardiac dysfunction. To address this mechanism, we synthesized the 1-DASA-ATPMS@CP1@Eth nanocomposite and first quantitatively evaluated its peroxidase-like activity using the TMB/H₂O₂ system (Figure 4). In the colorimetric assay, both CP1 and 1-DASA-ATPMS@CP1 exhibited a baseline absorbance peak at 652 nm, whereas Eth-loaded 1-DASA-ATPMS@CP1@Eth showed a significantly higher absorbance, indicating that Eth incorporation markedly enhances the nanozyme-like catalytic activity (Figure 4a). Subsequently, by varying the concentrations of TMB or H₂O₂ and applying Michaelis–Menten kinetics (via Lineweaver–Burk fitting: 1/V = (K_m/Vmax)(1/[S]) + 1/V_max), we found that 1-DASA-ATPMS@CP1@Eth has a much lower Km compared to CP1 and 1-DASA-ATPMS@CP1, demonstrating a higher substrate affinity and superior catalytic efficiency (Figure 4b). To elucidate the catalytic mechanism, radical scavenging experiments were performed using DMPO as a spin trap and monitored by electron spin resonance (ESR). As shown in Figure 4(c), 1-DASA-ATPMS@CP1@Eth generated typical four-line signals at a 1:2:2:1 ratio corresponding to ·OH, while Figure 4(d) shows the characteristic 1:1:1:1 quartet for ·O₂⁻; in both cases, the signal intensities were significantly greater than those of CP1 and 1-DASA-ATPMS@CP1, indicating that Eth loading amplifies the generation of highly oxidizing radicals. Furthermore, CV curves (Figure 4e) in the presence of TMB display pronounced redox peaks, confirming that the Dy³⁺/Dy²⁺ redox couple within the nanocomposite effectively promotes TMB oxidation. In addition to catalytic activity, the physicochemical properties of the nanocomposite were systematically evaluated. Dynamic light scattering (DLS) analysis (Figure 4f) revealed that 1-DASA-ATPMS@CP1@Eth possessed a relatively uniform particle size distribution centered at ∼772 nm, suggesting good dispersion stability. Zeta potential measurements (Figure 4g) further indicated a surface charge of approximately –25 mV, reflecting sufficient electrostatic repulsion to prevent aggregation. These results collectively confirm that 1-DASA-ATPMS@CP1@Eth not only exhibits enhanced catalytic efficiency but also maintains favorable colloidal stability, ensuring its applicability in biological environments.

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Figure 4.

(a) UV–vis spectra of CP1, 1-DASA-ATPMS@CP1, and 1-DASA-ATPMS@CP1@Eth in TMB/H₂O₂/NaAc–HAc (pH 4.0) (inset: color changes). (b) Impact of radical scavengers on peroxidase-ESR-like activity. (c) ESR of DMPO–·OH for CP1, 1-DASA-ATPMS@CP1, and 1-DASA-ATPMS@CP1@Eth. (d) ESR of DMPO–·O₂⁻ for the same samples. (e) CV curves, (f) DLS size; (g) Zeta potential.

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3.5. Fluorescence and electrochemical performance of 1-DASA-ATPMS@CP1@Eth

As shown in Figure 5(a), 1-DASA-ATPMS@CP1 exhibits a maximum excitation peak at approximately 330 nm and a bright blue emission centered around 460 nm. This fluorescence emission is attributable to the conjugated structure of the CP1 ligand, which absorbs photons to promote electrons to higher energy levels and subsequently emits light as they relax to lower energy states. After loading Eth into 1-DASA-ATPMS@CP1 (yielding 1-DASA-ATPMS@CP1@Eth), the emission peak remains at ∼460 nm, but its intensity is slightly reduced (Figure 5b), likely due to partial quenching or competitive absorption effects of Eth molecules on the CP1 fluorophore.

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Figure 5.

(a) Excitation and emission spectra of 1-DASA-ATPMS@CP1@Eth, (b) Fluorescence spectra of 1-DASA-ATPMS@CP1 and 1-DASA-ATPMS@CP1@Eth. (c) CV responses of the different materials. (d) EIS responses of the different materials.

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Figure 5(c) presents the CV curves for 1-DASA-ATPMS@CP1@Eth. Compared to the bare CP1-modified electrode, both 1-DASA-ATPMS@CP1 and 1-DASA-ATPMS@CP1@Eth electrodes show significantly increased peak currents, with the 1-DASA-ATPMS@CP1@Eth electrode displaying a peak current approximately 1.7 times that of the bare CP1 electrode. This enhancement indicates that Eth loading not only increases the number of active sites on the electrode surface but also facilitates interfacial electron transfer via the intrinsic conductivity of Eth. Moreover, the EIS data in Figure 5(d) further corroborate this finding: 1-DASA-ATPMS@CP1@Eth exhibits the lowest charge-transfer resistance (Rct≈70 Ω), markedly lower than that of 1-DASA-ATPMS@CP1 (≈ 90 Ω) and bare CP1 (≈ 120 Ω). Collectively, these results confirm that Eth loading optimizes both electron transport and conductivity of 1-DASA-ATPMS@CP1@Eth, laying a solid foundation for subsequent electrochemical sensing and catalytic applications.

3.6. Analytical performance of the proposed aptasensor

To systematically evaluate the trimodal sensing performance, we measured the UV–visible absorption, fluorescence emission, and electrochemical responses of Ang II under optimized conditions. All calibration curves were obtained from three independent measurements (n = 3), and the corresponding error bars have been shown in Figure 6. The raw calibration plots (prior to logarithmic transformation) are provided in the Supporting Information. Limits of detection (LOD) were calculated using the standard formula LOD = 3σ/k, where σ represents the standard deviation of the blank measurements and k is the slope obtained from linear regression. In the colorimetric mode, Ang II concentrations from 0.5 to 200 ng mL^-1 yielded a progressive increase in absorbance at 650 nm (Figure 6a). The calibration plot displayed an excellent linear relationship with log C_AngII (R² = 0.9961), and the fitted regression equation was y = 0.2337x + 0.4618 (Figure 6b), giving a slope k = 0.2337. Based on triple blank measurements (σ = 0.0123), the calculated LOD was: LOD=3σ/k=(3×0.0123)/0.2337=0.158 ng mL^-1, which agrees with the experimentally determined value of 0.16 ng mL^-1. In the fluorescence mode (Figure 6c), Ang II induced a concentration-dependent enhancement in emission intensity at 460 nm. The calibration curve (Figure 6d) exhibited strong linearity over the range 1–800 ng mL^-1, with the regression equation y = 930.3722x + 2856.7372 and R² = 0.9942, giving a slope k = 930.3722. Using the blank standard deviation (σ = 235.8), the LOD was determined as:

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Figure 6.

(a) UV–Vis absorption spectra at various Ang II concentrations; (b) Calibration curve of relative absorbance; (c) Fluorescence emission spectra at various; (d) Calibration curve of relative fluorescence intensity; (e) EIS spectra of the nanocomposite-modified electrode; (f) Linear fit of ΔRct vs. log[Ang II].

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LOD=(3×235.893)/0.3722=0.76 ng mL^-1, consistent with the observed sensitivity. In the electrochemical mode (Figure 6e), Ang II binding produced increasing charge-transfer resistance (Rct). The logarithmic calibration plot (Figure 6f) showed excellent linearity (R² = 0.9949) with the regression equation y = 73.4718x + 388.0328, giving a slope k = 73.4718. With a blank deviation of σ = 0.0213 Ω, the LOD was calculated as: LOD=(3×0.0213)/73.4718=8.7×10⁻⁴ ng mL^-1=0.87 pg mL^-1 demonstrating the superior sensitivity of the electrochemical pathway. Together, these results confirm that the 1-DASA-ATPMS@CP1@Eth sensing platform provides highly reliable and reproducible trimodal detection, supported by rigorous statistical analysis, transparent LOD calculations, and well-defined calibration behaviors.

3.7. In vitro release profile of 1-DASA-ATPMS@CP1@Eth

Figure 7(a) shows the in vitro release behavior of 1-DASA-ATPMS@CP1@Eth under different pH conditions, demonstrating a pronounced pH-responsive property. In an acidic environment (pH = 5.0), drug release was the most rapid, reaching approximately 65% within 10 h and nearly 90% after 96 h. Under weakly acidic conditions (pH = 6.4), the release rate was relatively moderate, with a final release of about 80%. In contrast, at physiological pH (7.4), drug release was much slower, with only about 60% released after 96 h. These findings indicate that the system enables accelerated release in acidic pathological microenvironments, while minimizing premature leakage under normal physiological conditions, thereby improving targeting and safety. Fluorescence stability tests in Figure 7(b) further confirmed that the fluorescence intensity of 1-DASA-ATPMS@CP1@Eth remained above 95% throughout 96 h, with negligible decay, demonstrating excellent photostability and structural integrity in vitro. Collectively, 1-DASA-ATPMS@CP1@Eth exhibits both pH-responsive targeted drug release and high stability, highlighting its promising potential for the treatment of osteoporosis-related diseases.

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Figure 7.

(a) In vitro drug release profile of 1-DASA-ATPMS@CP1@Eth under different pH conditions (7.4, 6.4, and 5.0). (b) Fluorescence stability of 1-DASA-ATPMS@CP1@Eth over 96 h. (c) Time-resolved fluorescence decay curves of 1-DASA-ATPMS@CP1 and 1-DASA-ATPMS@CP1@Eth.

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Importantly, the fluorescence lifetime decay profiles obtained by TCSPC (Figure 7c) further elucidate the excited-state dynamics of the system. The pristine 1-DASA-ATPMS@CP1 exhibits a relatively long fluorescence lifetime, consistent with its intrinsic relaxation behavior. Upon Eth incorporation, the composite 1-DASA-ATPMS@CP1@Eth shows a distinctly shortened lifetime, indicating the emergence of additional non-radiative decay pathways or photoinduced energy-transfer interactions between Eth and the CP1-based chromophore. Quantitative photophysical measurements support this interpretation: the absolute quantum yield (Φ) of 1-DASA-ATPMS@CP1@Eth decreases from Φ = 0.32 to Φ = 0.18 after binding Ang II, while time-resolved fluorescence analysis reveals a biexponential decay with lifetimes of τ₁ = 3.25 ns and τ₂ = 1.12 ns. These shortened lifetimes and reduced quantum yields confirm that Eth loading alters the local electronic environment within the nanocomposite, facilitating analyte-responsive modulation of radiative and non-radiative decay channels. Such behavior is fully consistent with the slight fluorescence quenching observed in the steady-state spectra and provides direct evidence of intermolecular interactions between Eth and the DASA-ATPMS@CP1 host matrix.

3.8. Kinetic modeling of drug release

The drug release kinetics were analyzed using several classical models to elucidate the underlying release mechanism (Figure 8). The zero-order kinetic model (Figure 8a) showed a linear increase of cumulative release with time, but with a relatively low correlation coefficient (R² = 0.6443), suggesting that the process is not purely time-dependent. In contrast, the first-order model (Figure 8b) exhibited a significantly better fit (R² = 0.9513), indicating that the release rate is strongly related to the remaining drug concentration. The Higuchi model (Figure 8c) also demonstrated a good correlation (R² = 0.8708), highlighting the contribution of diffusion during the release process. Notably, the Korsmeyer–Peppas model (Figure 8d) provided the best fit, with an excellent correlation coefficient (R² = 0.9940), suggesting that the release follows a non-Fickian diffusion profile driven by the combined effects of diffusion and carrier degradation.

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Figure 8.

Drug release kinetics of 1-DASA-ATPMS@CP1@Eth fitted to (a) zero-order, (b) first-order, (c) Higuchi, and (d) Korsmeyer–Peppas models.

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Taken together, these results indicate that the release of 1-DASA-ATPMS@CP1@Eth is predominantly governed by a synergistic mechanism involving polymer degradation and diffusion. Under acidic conditions (simulating pathological environments), the system exhibited accelerated release, while maintaining a relatively sustained release profile under neutral pH. Such environment-responsive behavior favors efficient drug delivery at lesion sites while minimizing premature release in healthy tissues, thereby enhancing both therapeutic specificity and safety.

3.9. 1-DASA-ATPMS@CP1@Eth inhibited Ang II-induced apoptosis in cardiomyocytes

Cardiomyocyte viability assessed by CCK-8 assay showed that Ang II (1 μM) markedly reduced survival (p<0.0001), confirming successful apoptosis induction (Figure 9a). Treatment with 40 μg/mL 1-DASA-ATPMS@CP1@Eth restored survival to 92.1%, significantly higher than both the model (p<0.001) and free drug groups (75.9%, p<0.01), while blank nanoparticles had no effect (p>0.05). At the molecular level, Ang II strongly up-regulated Bax mRNA (p<0.0001) (Figure 9b), which was suppressed to near-normal levels by nanoparticle treatment (p<0.0001); free drug also reduced Bax but less effectively (p<0.0001). These results demonstrate that 1-DASA-ATPMS@CP1@Eth exerts potent anti-apoptotic effects, surpassing the free drug. Importantly, its biocompatibility, targeted delivery, and efficacy highlight promise for in vivo applications. Future studies will explore its therapeutic potential in hypertensive rodent models by evaluating cardiac function, injury biomarkers, apoptosis-related gene expression, and carrier biodistribution, with parallel comparison to free drug to validate translational advantages.

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Figure 9.

(a) Cell viability of cardiomyocytes in each group detected by CCK-8 assay. (b) Relative expression of Bax mRNA in each group was detected by qPCR. Data are presented as mean ± SD from three independent experiments (n = 3). **, p < 0.01; ***, p < 0.001; ****, p < 0.0001.

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We developed a multifunctional 1-DASA-ATPMS@CP1@Eth nanocomposite capable of integrating colorimetric, fluorescence, and electrochemical responses with therapeutic intervention [27]. The mechanistic basis of its trimodal behavior arises from the synergistic interaction between the porous CP1 nanozyme core, the DASA-ATPMS functional shell, and the encapsulated Eth. Control experiments using CP1 alone or DASA-ATPMS@CP1 (without Eth) confirmed negligible optical and electrochemical responses toward Ang II, demonstrating that the enhanced sensing and therapeutic effects originate from the cooperative architecture of the hybrid rather than its isolated components [28]. Mechanistically, Ang II binding induces pronounced perturbations in the photophysical environment of the composite. Fluorescence lifetime analysis revealed a transition from the longer decay profile of pristine DASA-ATPMS@CP1 to a biexponential decay (τ₁ = 3.25 ns, τ₂ = 1.12 ns) after Eth incorporation, together with a decrease in quantum yield (Φ = 0.32→0.18 upon Ang II exposure). These changes indicate the emergence of non-radiative decay pathways facilitated by energy-transfer or dipole interactions between Eth and the DASA–CP1 matrix [29]. Concurrently, Ang II adsorption imposes interfacial resistance on the porous CP1 framework, increasing R_ct and modulating electron-transfer behavior, which correlates well with the colorimetric and fluorescence responses [30]. Therapeutically, Eth-loaded nanoparticles significantly attenuated Ang II–induced cytotoxicity compared to free Eth, consistent with improved intracellular delivery, enhanced stability, and microenvironment-responsive release enabled by the DASA-ATPMS shell [31]. The CP1 core further contributes peroxidase-like catalytic activity, allowing the nanocomposite to modulate oxidative stress, one of the central pathological drivers associated with Ang II signaling [32]. Together, these findings support a unified mechanistic model in which (i) Eth-mediated modulation of the local electronic environment, (ii) Ang II–induced alteration of CP1 electron-transfer channels, and (iii) DASA-assisted optical switching collectively give rise to the composite’s trimodal sensing behavior and enhanced therapeutic performance. While the present study elucidates the principal mechanisms and validates the platform in vitro, future work involving broader apoptotic markers and in vivo hypertensive models will be essential to fully establish its diagnostic and therapeutic potential.