English

• Nanomedicine offers significant potential for cancer therapy through the use of nanoscale materials, including liposomes, micelles, and various nanoparticles. These nanomaterials leverage the enhanced permeability and retention (EPR) effect to preferentially accumulate in tumor tissues, thereby improving therapeutic efficacy while minimizing off-target effects [1-10]. Nonetheless, < 1% of nanomedicines achieve effective delivery to solid tumors [11]. Self-assembled nanomedicines, stabilized by non-covalent intermolecular forces, often face premature drug leakage and rapid bloodstream clearance due to serum protein adsorption, which triggers their removal by macrophages in the reticuloendothelial system (RES) after intravenous administration [12-16]. The surface charge of nanoparticles plays a crucial role in determining their in vivo behaviors [17-19]. Positively charged nanoparticles demonstrate increased internalization by tumor cells but suffer from rapid RES clearance, which reduces their delivery efficacy [20, 21]. In contrast, negatively charged nanoparticles are frequently used to evade RES clearance, yet their limited penetration ability results in inadequate cellular uptake by tumor cells [22-28]. Therefore, balancing the stability of nanomedicines with their efficient cellular uptake within the tumor microenvironment (TME) remains a major challenge.

  To address these issues, charge reversal strategies have emerged as novel approach [29-31]. By dynamically modulating the surface charge of nanoparticles in response to specific intra- or extracellular signals, such as pH fluctuations, glutathione (GSH) levels, reactive oxygen species, and enzymes, charge reversal can facilitate an optimal balance between evading RES clearance and enhancing cellular uptake within the TME [32-37]. Despite these promising advances, significant challenges remain, particularly concerning the long-term stability of charge-reversible nanomedicines [38-41]. Instability often results in premature drug leakage, leading to non-specific toxicity and reduced therapeutic efficacy [42]. Nanogels, a class of nanosized hydrogels, are characterized by a three-dimensional cross-linked structure that provides exceptional stability and a high capacity for surface modification [43]. Compared to conventional nanomedicines, nanogels benefit from their high hydrophilicity and polymer-based hydrogel structure, which facilitate enhanced membrane deformation and, consequently, promote more efficient cellular internalization [44-46]. Surface functionalization of nanogels, particularly via charge-reversal strategies, enhances their stability, targeting efficiency, and cellular uptake, thereby improving therapeutic outcomes and reducing off-target effects.

  Herein, we developed a novel charge-reversible nanogel for targeted paclitaxel (PTX) dimers (DPP) delivery, designed to enhance stability and precision within the TME. This nanocarrier, with a dynamically responsive surface charge, optimizes drug delivery efficacy while minimizing systemic toxicity. DPP was encapsulated via free radical polymerization, significantly improving its solubility and stability. Upon reaching the tumor site, the charge-reversal mechanism promotes cellular uptake and localized drug release, enhancing therapeutic efficacy. In the high-GSH tumor cells, DPP releases active PTX through disulfide bond cleavage, reducing toxicity and ensuring targeted action. Experimental results showed marked improvements in delivery efficiency and reduced off-target effects, underscoring the potential of this approach for cancer therapy. Initially, DPP was synthesized from PTX and a disulfide-containing aliphatic dicarboxylic acid, using 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC·HCl) and 4-dimethylaminopyridine (DMAP) as catalysts (Fig. S1 in Supporting information), with synthesis confirmed by proton nuclear magnetic resonance and mass spectrometry (Figs. S2 and S3 in Supporting information). As shown in Scheme 1, nanogels were prepared by solvent exchange with F127, forming DPP@F127. Further free radical polymerization with acrylamide (AAM), N-(3-aminopropyl)-methacrylamide (APM), and disulfide crosslinker N, N′-bis(acryloyl)cystamine (BAC) produced n(DPP@F127). To enable pH-triggered charge reversal, 2,3-dimethylmaleic anhydride (DMA) was conjugated to amino groups on n(DPP@F127), forming n(DPP@F127)_R with negatively charged carboxyl groups at neutral and alkaline pH that convert to positively charged amines under acidic condition. Furthermore, the nanogels escape lysosomes and release PTX in response to high intracellular GSH, enhancing cytotoxicity against tumor cells.

  Scheme 1

  

  Scheme 1.  Schematic illustration of charge-reversal dimeric PTX prodrug nanogels for tumor treatment.

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  As shown in Fig. S4 (Supporting information), the infrared spectra exhibit a disulfide bond vibration absorption peak at 540 and 637 cm⁻¹, indicating that the stable disulfide bond structure is preserved in DPP@F127, n(DPP@F127), and n(DPP@F127)_R before and after crosslinking. Moreover, the disappearance of the crystalline peaks of n(DPP@F127) and n(DPP@F127)_R in the XRD patterns (Fig. S5 in Supporting information) suggests that the nanogel transitions to an amorphous state following crosslinking modification. This further confirms the successful crosslinking reaction.

  Transmission electron microscopy (TEM) images (Fig. 1a) and dynamic light scattering (DLS) analysis (Fig. 1b) reveal that DPP@F127, n(DPP@F127), and n(DPP@F127)_R are spherical with hydrodynamic diameters of 35.32, 94.01, and 117.4 nm, respectively, in clear solutions. The increased particle sizes of n(DPP@F127) and n(DPP@F127)_R confirm successful polymerization. To assess stability, we monitored the particle size and polydispersity index (PDI) of DPP@F127, n(DPP@F127), and n(DPP@F127)_R in phosphate-buffered saline (PBS, pH 7.4) and Dulbecco's modified Eagle medium (DMEM), as well as in PBS containing 10% fetal bovine serum (FBS). As shown in Figs. S6 and S7 (Supporting information), the particle size and PDI of all nanoparticles remained largely unchanged in PBS and DMEM. In contrast, in PBS containing FBS, DPP@F127 exhibited significant size fluctuations, increase in particle size and PDI, and substantial flocculent precipitation appeared by day 5, indicating increased heterogeneity and instability due to protein adsorption (Fig. 1c). The n(DPP@F127) and n(DPP@F127)_R maintained stable sizes and PDI values throughout the experiment (Fig. 1d and e), reflecting robust colloidal stability attributed to the crosslinked network's resistance to protein adsorption. This stability preserves structural integrity, minimizing premature drug release and aggregation, and supports consistent drug delivery in systemic circulation.

  Figure 1

  

  Figure 1.  Characterization and charge-reversal capacity. (a) TEM image and (b) sizes of (Ⅰ) DPP@F127, (Ⅱ) n(DPP@F127), and (Ⅲ) n(DPP@F127)_R. Scale bars, 100 nm. The change of size and size distribution of (c) DPP@F127, (d) n(DPP@F127) and (e) n(DPP@F127)_R for 7 days. (f) Zeta potential change of n(DPP@F127)_R at different pH PBS buffer. (g) Size change in n(DPP@F127) with different DTT concentrations. (h) The DPP release behavior of DPP@F127 and n(DPP@F127) with or without DTT, as determined by HPLC. Data are expressed as the mean ± SD (n = 3). **P < 0.01, ****P < 0.0001.

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  To evaluate the charge-reversal capability of n(DPP@F127)_R, we measured its zeta potential in PBS across various pH levels over 24 h (Fig. 1f). At pH 7.4, the zeta potential remained stable, showing no charge modulation. In acidic conditions, however, n(DPP@F127)_R exhibited significant charge reversal, shifting from −6.72 mV to 3.36 mV at pH 6.5 and from −6.41 mV to 7.44 mV at pH 5.0, confirming its effective charge-reversal capability. Further testing showed that both DPP@F127 and n(DPP@F127) consistently maintained their charge at pH 6.5 and 7.4 (Fig. S8 in Supporting information). Notably, n(DPP@F127)_R exhibited pH-sensitive properties, remaining negatively charged at pH 7.4 and shifting to a positive charge at pH 6.5, highlighting its potential for targeted drug delivery in the TME. Additionally, as shown in Fig. S9 (Supporting information), we examined particle size changes of DPP@F127, n(DPP@F127), and n(DPP@F127)_R under acidic conditions. While all nanoparticles increased in size with decreasing pH, the crosslinked n(DPP@F127) and n(DPP@F127)_R exhibited relatively minor size changes, indicating enhanced stability from crosslinking.

  We evaluated the drug release of DPP@F127, n(DPP@F127) and n(DPP@F127)_R by monitoring particle size changes in the presence of dithiothreitol (DTT), a reducing agent analogous to GSH (Fig. 1g). Particle size increased with higher DTT concentrations, as DTT cleaved disulfide bonds within the crosslinked structure, relaxing the nanogels. This reduction-triggered release mechanism was confirmed by high-performance liquid chromatography (HPLC), monitored at 231 nm (Fig. S10 in Supporting information). The drug loading contents of DPP@F127, n(DPP@F127), and n(DPP@F127)_R were 16.87% ± 0.19%, 14.86% ± 0.11%, and 14.25% ± 0.14%, respectively, as shown in Table S1 (Supporting information). As shown in Fig. 1h, n(DPP@F127) and n(DPP@F127)_R released approximately 60% of PTX within 30 h under DTT conditions. In contrast, without DTT, only 20% PTX release occurred, underscoring the DTT-sensitive nature of these nanogels. We conducted repeatability validation using 15 µmol/L GSH, and a similar trend was observed in Fig. S11 (Supporting information). This substantial release under reducing conditions suggests that high-GSH levels in tumor tissues could trigger localized PTX release, enhancing therapeutic efficacy while minimizing systemic toxicity. The charge-reversal and DTT-responsive properties of n(DPP@F127)_R offer significant potential for targeted cancer therapy. Charge reversal facilitates cellular uptake in the acidic TME, while reduction-responsive release enables precise drug delivery. Together, these features demonstrate the effectiveness of integrating charge-reversal and responsive release mechanisms into nanogel design for cancer treatment.

  Validating the delivery mechanism of the nanogels into cells is crucial for assessing their therapeutic potential. To investigate internalization, fluorescein isothiocyanate (FITC) was incorporated into nanogels using a similar construction process as for DPP. The resulting FITC-loaded nanogels, FITC@F127, n(FITC@F127), and n(FITC@F127)_R, were characterized for size and surface charge. As shown in Fig. S12 (Supporting information), these nanogels displayed spherical shapes with average hydrodynamic diameters of 34.26, 78.83, and 91.56 nm, respectively. The zeta potentials of FITC@F127 and n(FITC@F127) were negative at both pH 7.4 and 6.5, whereas n(FITC@F127)_R exhibited notable charge reversal from negative to positive at pH 6.5, consistent with the behavior of n(DPP@F127)_R. These results indicate that n(FITC@F127)_R undergoes charge reversal in the acidic conditions typical of the TME (Fig. S13 in Supporting information). The fluorescence emission spectra revealed aggregation-induced quenching, suggesting that FITC was physically encapsulated within the nanogels via hydrophilic-hydrophobic interactions, rather than surface adsorption (Fig. S14 in Supporting information). This encapsulation model is essential for understanding nanogel degradation in the cellular environment.

  To ensure safety before cellular uptake experiments, we assessed L929 cell viability with the cell counting kit-8 (CCK-8) assay, treating cells with drug-free nanoparticles F127, n(F127), and n(F127)_R (0–100 µg/mL) at pH 7.4 and 6.5 for 24 h, showing minimal cytotoxicity and indicating low toxicity to normal fibroblasts (Fig. 2a). For cellular uptake assessment, MCF-7 cells were incubated with n(FITC@F127) and n(FITC@F127)_R at pH 7.4 and 6.5 for 6 h at 37 ℃. As shown in Fig. 2b, n(FITC@F127) exhibited minimal uptake differences between pH levels due to its consistent negative surface charge. In contrast, n(FITC@F127)_R displayed significantly enhanced uptake at pH 6.5, reflecting its increased positive charge under acidic conditions. Fluorescence quantification confirmed this, with n(FITC@F127)_R showing 11.6 times greater fluorescence intensity at pH 6.5 than at pH 7.4 (Fig. 2c and Fig. S15 in Supporting information). This substantial increase underscores the effectiveness of charge reversal in promoting cellular internalization. Next, n(FITC@F127) and n(FITC@F127)_R were incubated at 37 and 4 ℃ over a 6-h period. As shown in Fig. 2d, fluorescence was significantly lower at 4 ℃, indicating that endocytosis is temperature-dependent and likely mediated by energy-driven mechanisms, such as clathrin- or receptor-mediated pathways. The data in Fig. 2e indicate that at both pH 7.4 and 6.5, the green fluorescence from n(FITC@F127) co-localizes with Lyso-Tracker red, with colocalization percentages of 51% and 75%, respectively, which confirms lysosomal internalization through the endolysosomal pathway. For n(FITC@F127)_R, the fluorescence overlap with Lyso-Tracker at pH 6.5 (82%) suggests effective lysosomal targeting. However, the significantly reduced overlap at pH 7.4 (10%) points to potential issues related to nanogel degradation and increased membrane permeability, which might affect the targeting efficiency. These results highlight the role of pH-sensitive charge reversal in cellular uptake and intracellular trafficking.

  Figure 2

  

  Figure 2.  Cellular uptake. (a) Cell viability of F127, n(F127) and n(F127)_R. (b) Fluorescence microscopy images of MCF-7 cells incubated with n(FITC@F127) and n(FITC@F127)_R in PBS at pH 7.4 and 6.5. (c) Mean fluorescence intensity measured by FCM for n(FITC@F127)_R at pH 7.4 and 6.5. (d) Confocal laser scanning microscopy (CLSM) images of MCF-7 cells with n(FITC@F127) and n(FITC@F127)_R at 4 and 37 ℃ for 6 h. Nuclei and actin were stained with 4′, 6-diamidino-2-phenylindole, DAPI (blue) and DiI (red). (e) Lysosomal colocalization and semi-quantitative fluorescence analysis of n(FITC@F127) and n(FITC@F127)_R at pH 7.4 and 6.5. Lysosome was stained with Lyso-Tracker (red), respectively. Scale bar: 100 µm. ns, no significance. Data are presented as mean ± SD (n = 3). **P < 0.01, ***P < 0.001.

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  To assess the antiproliferative efficacy of DPP nanogels, we measured cell viability in MCF-7 and L929 cell lines using the CCK-8 assay, treating cells with DPP@F127, n(DPP@F127) and n(DPP@F127)_R at concentrations from 0 to 100 µg/mL at pH 7.4 and 6.5 for 24 h. Results indicated minimal cytotoxicity towards L929 cells under both pH conditions, demonstrating low toxicity to normal fibroblasts (Fig. S16 in Supporting information). In contrast, MCF-7 cells showed significant differences in viability based on formulation. As depicted in Fig. 3a, DPP@F127 had negligible effects on viability at either pH, indicating limited efficacy. Conversely, n(DPP@F127) reduced cell survival below 50% at 100 µg/mL under both pH conditions, correlating with increased DPP uptake and enhanced cytotoxicity (Fig. 3b). Notably, n(DPP@F127)_R reduced MCF-7 viability to 43% at pH 6.5 and 79% at pH 7.4 (Fig. 3c), likely due to its surface charge dynamics. As shown in Fig. 3d, live-dead staining revealed that n(DPP@F127)_R exhibited significantly more red fluorescence at pH 6.5 than at 7.4, while n(DPP@F127) induced extensive apoptosis at both pH levels. Flow cytometry results showed a similar trend, with n(DPP@F127) and n(DPP@F127)_R inducing over 50% late apoptosis in cells, compared to only 7% apoptosis observed with DPP@F127 (Fig. S17 in Supporting information). The consistently negative DPP@F127 experiences electrostatic repulsion from the cell membrane, limiting uptake, while positively charged n(DPP@F127) is more readily internalized. In high-GSH environments typical of tumor cells, PTX is efficiently released, targeting cancer cells while sparing normal cells. Additionally, n(DPP@F127)_R undergoes charge reversal in acidic conditions, acquiring a positive charge at pH 6.5, which enhances cellular uptake and drug delivery within the TME.

  Figure 3

  

  Figure 3.  Cellular cytotoxicity. Cytotoxicity of (a) DPP@F127, (b) n(DPP@F127), and (c) n(DPP@F127)_R toward MCF-7 cells at pH 7.4 and 6.5 conditions via CCK-8 assays. Data are presented as mean ± SD (n = 6). (d) Fluorescence images of calcein-AM/PI-co-stained MCF-7 cells (green for live cells and red for dead cells) incubated with PBS, DPP@F127, n(DPP@F127) and n(DPP@F127)_R at DPP concentration of 100 µg/mL for 24 h. Scale bar: 100 µm.

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  Accurate drug efficacy assessment necessitates in vivo models to complement in vitro data for a comprehensive antitumor evaluation. Hemolysis assays demonstrated that DPP@F127, n(DPP@F127), and n(DPP@F127)_R exhibit excellent biocompatibility, with hemolysis rates consistently below 5%, highlighting their potential for safe biological applications (Fig. S18 in Supporting information). All animal experiments have been approved (No. 2023–0132) by the Animal Welfare and Ethics Committee of Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, and carried out according to the NIH guidelines for the care and use of laboratory animals (NIH publication No. 85–23 Rev. 1985). Female BALB/c mice were obtained and raised under required conditions. We conducted a pharmacokinetic study of DPP@F127, n(DPP@F127) and n(DPP@F127)_R in BALB/c mice (18–22 g) at a dose of 15 mg/kg, collecting blood samples at specific intervals to analyze serum DPP concentrations by HPLC. As shown in Fig. 4a, n(DPP@F127) exhibited a plasma elimination half-life of 1.65 h, four times that of DPP@F127 (0.42 h). Notably, n(DPP@F127)_R achieved a half-life of 5.67 h, indicating significantly enhanced circulatory stability.

  Figure 4

  

  Figure 4.  In vivo antitumor efficacy of DPP@F127, n(DPP@F127) and n(DPP@F127)_R. (a) Pharmacokinetic profiles of DPP@F127, n(DPP@F127), and n(DPP@F127)_R following systemic administration; bars represent SD (n = 3). (b) Schematic of animal treatment administration: 15 mg/kg equivalent DPP dose for saline, DPP@F127, n(DPP@F127) and n(DPP@F127)_R groups. (c, d) Tumor growth curves, (e) tumor weights, and (f) representative images of excised tumors from treatment groups; error bars represent SD (n = 6). (g) H & E staining of tumor sections from saline, DPP@F127, n(DPP@F127), and n(DPP@F127)_R groups. Scale bar: 100 µm. **P < 0.01, ***P < 0.001, ****P < 0.0001.

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  In vivo antitumor efficacy was further evaluated in a 4T1 breast cancer xenograft BALB/c mouse model, with mice randomly assigned to treatment groups: saline, DPP@F127, n(DPP@F127), and n(DPP@F127)_R (Fig. 4b). After tumors reached ~50 mm³, mice received intravenous doses of 15 mg/kg every two days for a total of five treatments, and were sacrificed on day 14. As illustrated in Fig. 4c and d, DPP@F127, n(DPP@F127) and n(DPP@F127)_R suppressed tumor growth, with n(DPP@F127)_R demonstrating the most substantial inhibition. This was confirmed by reduced tumor weights and photographs (Fig. 4e and f). Histological analysis using hematoxylin and eosin (H & E) staining revealed significant nuclear pyknosis and extensive tumor cell ablation in n(DPP@F127)_R-treated samples, compared to those treated with DPP@F127 and n(DPP@F127) (Fig. 4g).

  In addition to evaluating the therapeutic efficacy of DPP@F127, n(DPP@F127) and n(DPP@F127)_R, we assessed their systemic toxicity to ensure a suitable safety profile for potential clinical application. Mice treated with various nanogel formulations exhibited no significant changes in body weight, which remained within normal physiological ranges throughout the study (Fig. S19 in Supporting information). Histological analyses were conducted on key organs, revealed no adverse histopathological changes in these organs in mice treated with the DPP@F127, n(DPP@F127) and n(DPP@F127)_R (Fig. 5a). Serum biochemical analyses were performed to assess kidney and liver function as well as other blood parameters. The results showed no significant differences between mice treated with DPP@F127, n(DPP@F127) and n(DPP@F127)_R and untreated controls, suggesting that the nanogels did not adversely affect systemic organ function (Fig. 5b). The minimal systemic toxicity and effective antitumor activity of charge-reversal PTX nanogels are promising candidates for breast cancer therapy.

  Figure 5

  

  Figure 5.  In vivo safety evaluation: (a) Histological assessment using H & E staining of various organs (heart, liver, spleen, lung, and kidney) in mice. Scale bar: 100 µm. (b) Serum biochemical analysis of blood parameters and kidney and liver function indicators. HCT, hematocrit; HGB, hemoglobin; MCV, mean corpuscular volume; WBC, white blood cell; RBC, red blood cell; PLT, platelet; UREA, urea; CREA, creatinine; ALT, alanine aminotransferase; AST, aspartate transaminase. Data are presented as mean ± SD (n = 6).

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  In summary, this study presents a charge-reversible nanocarrier strategy for PTX delivery in breast cancer therapy, achieving enhanced stability, targeted drug delivery, and improved therapeutic outcomes. The charge-reversal mechanism enables surface charge modulation in response to the acidic TME, optimizing cellular uptake and localized drug release. This approach ensures effective PTX release in high-GSH conditions typical of cancer cells, reducing off-target toxicity while enhancing therapeutic efficacy. In vivo studies confirmed the superiority of n(DPP@F127)_R, which demonstrated an extended plasma half-life and superior tumor suppression compared to other formulations. Notably, these nanocarriers exhibited excellent biosafety with no significant systemic toxicity. Overall, the integration of charge reversal and responsive release in this nanocarrier design represents a promising advancement in targeted cancer therapy, enhancing the specificity and effectiveness of chemotherapy.

  Declaration of competing interest

  The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

  CRediT authorship contribution statement

  Haotian Shi: Writing – original draft, Resources, Investigation. Yuchao Luo: Resources, Investigation, Data curation. Song Zhang: Visualization, Data curation. Meijun Zhao: Investigation. Chaoyong Liu: Visualization, Project administration. Qing Pei: Resources, Formal analysis. Helei Wang: Writing – review & editing, Validation. Qiong Dai: Project administration. Zhigang Xie: Validation, Supervision, Conceptualization. Bin Xu: Writing – review & editing, Supervision, Project administration, Funding acquisition. Wenjing Tian: Writing – review & editing, Supervision, Resources, Funding acquisition.

  Acknowledgments

  This work was supported by the Natural Science Foundation of Jilin Province (No. 20240101003JJ), the National Natural Science Foundation of China (Nos. 22275065, 52073116).

  Supplementary materials

  Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.cclet.2024.110775.

  
  
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  Scheme 1  Schematic illustration of charge-reversal dimeric PTX prodrug nanogels for tumor treatment.

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  Figure 1  Characterization and charge-reversal capacity. (a) TEM image and (b) sizes of (Ⅰ) DPP@F127, (Ⅱ) n(DPP@F127), and (Ⅲ) n(DPP@F127)_R. Scale bars, 100 nm. The change of size and size distribution of (c) DPP@F127, (d) n(DPP@F127) and (e) n(DPP@F127)_R for 7 days. (f) Zeta potential change of n(DPP@F127)_R at different pH PBS buffer. (g) Size change in n(DPP@F127) with different DTT concentrations. (h) The DPP release behavior of DPP@F127 and n(DPP@F127) with or without DTT, as determined by HPLC. Data are expressed as the mean ± SD (n = 3). **P < 0.01, ****P < 0.0001.

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  Figure 2  Cellular uptake. (a) Cell viability of F127, n(F127) and n(F127)_R. (b) Fluorescence microscopy images of MCF-7 cells incubated with n(FITC@F127) and n(FITC@F127)_R in PBS at pH 7.4 and 6.5. (c) Mean fluorescence intensity measured by FCM for n(FITC@F127)_R at pH 7.4 and 6.5. (d) Confocal laser scanning microscopy (CLSM) images of MCF-7 cells with n(FITC@F127) and n(FITC@F127)_R at 4 and 37 ℃ for 6 h. Nuclei and actin were stained with 4′, 6-diamidino-2-phenylindole, DAPI (blue) and DiI (red). (e) Lysosomal colocalization and semi-quantitative fluorescence analysis of n(FITC@F127) and n(FITC@F127)_R at pH 7.4 and 6.5. Lysosome was stained with Lyso-Tracker (red), respectively. Scale bar: 100 µm. ns, no significance. Data are presented as mean ± SD (n = 3). **P < 0.01, ***P < 0.001.

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  Figure 3  Cellular cytotoxicity. Cytotoxicity of (a) DPP@F127, (b) n(DPP@F127), and (c) n(DPP@F127)_R toward MCF-7 cells at pH 7.4 and 6.5 conditions via CCK-8 assays. Data are presented as mean ± SD (n = 6). (d) Fluorescence images of calcein-AM/PI-co-stained MCF-7 cells (green for live cells and red for dead cells) incubated with PBS, DPP@F127, n(DPP@F127) and n(DPP@F127)_R at DPP concentration of 100 µg/mL for 24 h. Scale bar: 100 µm.

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  Figure 4  In vivo antitumor efficacy of DPP@F127, n(DPP@F127) and n(DPP@F127)_R. (a) Pharmacokinetic profiles of DPP@F127, n(DPP@F127), and n(DPP@F127)_R following systemic administration; bars represent SD (n = 3). (b) Schematic of animal treatment administration: 15 mg/kg equivalent DPP dose for saline, DPP@F127, n(DPP@F127) and n(DPP@F127)_R groups. (c, d) Tumor growth curves, (e) tumor weights, and (f) representative images of excised tumors from treatment groups; error bars represent SD (n = 6). (g) H & E staining of tumor sections from saline, DPP@F127, n(DPP@F127), and n(DPP@F127)_R groups. Scale bar: 100 µm. **P < 0.01, ***P < 0.001, ****P < 0.0001.

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  Figure 5  In vivo safety evaluation: (a) Histological assessment using H & E staining of various organs (heart, liver, spleen, lung, and kidney) in mice. Scale bar: 100 µm. (b) Serum biochemical analysis of blood parameters and kidney and liver function indicators. HCT, hematocrit; HGB, hemoglobin; MCV, mean corpuscular volume; WBC, white blood cell; RBC, red blood cell; PLT, platelet; UREA, urea; CREA, creatinine; ALT, alanine aminotransferase; AST, aspartate transaminase. Data are presented as mean ± SD (n = 6).

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