Charge-convertible MnO-coated liposomal nanocarriers boost doxorubicin delivery for potentiated breast cancer chemotherapy.

Introduction
Despite the rapid advancements in innovative cancer therapeutic modalities, chemotherapy remains the most widely used therapeutic approach in clinical oncology.1 However, the systemic administration of free chemotherapeutics often leads to suboptimal treatment efficacy and severe systemic toxicity, primarily due to their non-selective biodistribution and unfavorable pharmacokinetic profiles.2 The emergence of nanotechnology has enabled the development of nanoscale drug delivery systems, which aim to optimize drug pharmacokinetics and overcome these limitations.3
For nanomedicines to effectively deliver therapeutic payloads to the cytoplasm of cancer cells within solid tumors, they must traverse a sequential cascade of biological barriers, collectively termed the CAPIR cascade: circulation in the blood compartment (C), accumulation in the tumor (A), penetration into the tumor parenchyma (P), cellular internalization by tumor cells (I), and intracellular drug release (R).4 The design of nanocarriers capable of navigating the CAPIR cascade presents a significant challenge, as the functional requirements at each step often exhibit antagonistic characteristics. For instance, nanoparticles (NPs) leverage the enhanced permeability and retention (EPR) effect to achieve passive accumulation in tumors by evading renal excretion, yet, their large size and rigid structure impede efficient penetration through the dense extracellular matrix (ECM).5 Surface PEGylation improves NPs’ stealth properties and prolongs their circulation half-life, but it concurrently attenuates the cellular internalization efficiency due to reduced interactions with cell surface receptors.6 Additionally, negatively charged NPs demonstrate superior circulatory stability and reduced toxicity to normal tissues. In contrast, positively charged NPs exhibit enhanced cellular uptake in tumor tissues due to electrostatic interactions with the negatively charged cell membrane.7 To address these conflicting requirements, tumor-responsive smart nanomedicines with adaptive functional conversion have emerged as a promising strategy to enhance antitumor therapeutic activity.8,9
Beyond NPs-intrinsic properties, the tumor microenvironment (TME) poses additional barriers to effective cancer treatment.10 Tumor hypoxia, a hallmark of the TME, is closely associated with tumor progression and therapeutic resistance, and is characterized by the upregulation of hypoxia-inducible factor 1 (HIF-1α) in hypoxic niches.11 HIF-1α-driven transcription of connective tissue growth factor (CTGF) and collagen I lead to the formation of a dense ECM, which further restricts nanomedicine penetration into the tumor parenchyma and thereby compromise antitumor efficacy.12 In recent years, substantial research efforts have focused on modulating tumor hypoxia to improve treatment outcomes across various modalities, including chemotherapy,13 photodynamic therapy (PDT),14 and immunotherapy.15 A notable example is the development of manganese dioxide (MnO2)-based nanosystems, which can catalytically decompose endogenous hydrogen peroxide (H2O2) in the acidic TME to generate oxygen (Equation 1).16 Integrating tumor-responsive charge-conversion capabilities with hypoxia-modulating functions in a single nanoplatform and thus holds great promise for achieving synergistic enhancement of antitumor efficacy, addressing both the biophysical barriers of the CAPIR cascade and the biochemical challenges of the hypoxic TME.
Herein, we designed a MnO2-coated cationic liposomal nanocarrier to boost DOX delivery efficiency through multiple ways and finally potentiate chemotherapy (termed MnO2@CLDOX). As depicted in Scheme 1A, during the preparation of CaO2 NPs, DOX molecules were pre-loaded into the CaO2 matrix via chelation with Ca2+ ions.17 Subsequently, the resulting drug-loaded cores were stabilized using cationic phospholipid and PEGylated-phospholipid for improved druggability.18 Upon adding an aqueous solution of MnCl2, MnO2 NPs formed in situ on the liposomal surface via reaction with CaO2 (Equation 2). Notably, this in situ coating strategy integrates CaO2-mediated DOX loading and MnO2 shell formation within a single synthetic step, circumventing the inherent complexity of post-synthesis modification and undesirable drug leakage associated with traditional multi-step fabrication processes. More importantly, this work represents the first reported approach to couple electronegative MnO2 materials with cationic liposomes to enable TME-triggered charge reversal—a design that transcends the conventional application of MnO2. By integrating the intrinsic acid/H2O2 dual responsiveness, oxygen-generating capacity, and electronegativity of MnO2 with cationic liposome-mediated cellular internalization, our MnO2@CLDOX system enables comprehensive optimization of each step in the CAPIR cascade, rather than relying on a single functional property of MnO2.
The underlying anticancer mechanisms are outlined as follows (Scheme 1B): (1) by EPR effects, the negatively charged MnO2@CLDOX passively accumulates in tumoral regions without causing severe side effects to normal tissues19; (2) the elevated H2O2 and acidic pH in TME disintegrate the MnO2 layers of NPs and uncover the drug-loaded liposomes CLDOX20; (3) the CLDOX exhibits enhanced cellular uptake via electrostatic interactions21; and (4) MnO2 dissolving reaction leads to O2 production, which further mediated HIF-1/CTGF/collagen I inhibition for boosting DOX tumor penetration. Through the rational design and construction of this smart/hypoxia-modulated nanocarrier, the CAPIR cascade of drug delivery was efficiently realized, thereby enhancing chemotherapy efficacy. We envision that these MnO2-coated liposomal nanocarriers also exhibit substantial potential in other O2-dependent cancer therapeutic modalities, including PDT and radiotherapy.

Results

Preparation and characterization
According to our previous study, CaO2 NPs were synthesized via a reaction involving CaCl2, H2O2, and NH3·H2O in a reverse microemulsion system.22 To fabricate drug-loaded CaO2 NPs encapsulated with a hydrophobic coating, DOX and DSPA were incorporated into the reverse microemulsion system. Under the alkaline conditions provided by NH3·H2O, DOX underwent deprotonation, enabling it to chelate with Ca2+ ions and form a Ca-DOX complex, subsequently entrapped within the CaO2 NPs.17 Likewise, the phosphate moieties of DSPA molecules coordinated with Ca2+ ions, facilitating the hydrophobic modification of the CaO2 NPs surface by assembling of long-chain alkyl groups.23 As depicted in Figure 1A, the purple-colored DSPA@DOX-CaO2 exhibited excellent solubility in non-polar solvents such as chloroform. From the image obtained by TEM, we found that the NPs’ size was about 50 nm with irregular morphology.
For further use, DSPE-PEG2000, cholesterol, and DOTAP were added to form CLDOX-CaO2 by a filming-rehydration method. To prepare MnO2-coated NPs, the obtained CLDOX-CaO2 NPs were added to an excess volume of MnCl2 solution. Highly water-soluble CaO2 NPs reacted rapidly with MnCl2 and produced MnO2 NPs that tend to deposit on the surface of cationic liposomes.24 After that, the solution color turned purple to brownish due to the formation of MnO2 NPs. As shown in Figure 1B, MnO2@CLDOX exhibits an increased particle size (about 150 nm) after coating with MnO2 and lipids. The hydrodynamic size of MnO2@CLDOX was measured to be 264 nm by DLS (Figure S1), sharply larger than the results obtained by TEM. This could be explained by the existence of PEG2000 and the hydration shell of NPs, which could not be observed by TEM. The Energy-dispersive X-ray spectroscopy (EDS) mapping recorded in Figure 1C demonstrated the existence of N, P and Mn, indicating the formation of lipids and MnO2 coatings. Moreover, no peak corresponding to the Ca element was detected in the EDS point scan of MnO2@CLDOX NPs (Figure S2). EDS line-scanning analysis demonstrates that MnO2@CLDOX exhibits a distinct core-shell structure, with MnO2 NPs localized in the outer liposomal layer. (Figure S3). The UV-Vis spectrum of MnO2@CLDOX showed a characteristic absorption band of DOX at 480 nm, which was also present in the free DOX spectrum (Figure 1D). The DOX loading content in MnO2@CLDOX was determined by HPLC to be 5.15 ± 0.10 wt %, while the Mn4+ loading capacity (2.9 ± 0.23 wt %) was measured by ICP-AAS.
CLDOX was prepared as a control group via the thin-film hydration method using the same phospholipid formulation. Meanwhile, MnO2@CL was manufactured as a blank carrier using the identical procedure to that for MnO2@CLDOX, except that DOX was omitted during preparation. Both nanomaterials exhibited particle sizes similar to MnO2@CLDOX (Figure S4). As depicted in Figure 1E, the zeta potential of CLDOX was measured to be 7.33 ± 1.22 mV, whereas that of MnO2@CL and MnO2@CLDOX exhibited a value of −12.7 ± 0.67 and −9.57 ± 0.96 mV, respectively. This significant shift of potential provides direct evidence for the successful coating of MnO2 on CLDOX.
To evaluate the suitability of MnO2@CLDOX as a nanomedicine, its stability was first assessed under simulated physiological conditions. As shown in Figure 1F, the average particle size of MnO2@CLDOX remained nearly unchanged (within the range of 220–280 nm) after incubation in saline and 10% FBS for one week. Moreover, the zeta potential of MnO2@CLDOX remained consistently negative in saline and 10% FBS (Figure 1G). These results indicated the good stability of MnO2@CLDOX under physiological conditions.

TME-responsive drug release and multifunctional evaluation
In contrast to normal cells, tumor tissues exhibit significantly elevated concentrations of H+ and H2O2.20,25 Accordingly, MnO2@CLDOX were expected to respond to TME in a H+ and H2O2-dependent manner. To validate this assumption, we first monitored the drug release profile of MnO2@CLDOX in solutions by the dynamic dialysis method. As exhibited in Figure 2A, under physiological conditions (pH 7.4), MnO2@CLDOX exhibited minimal drug leakage with a cumulative DOX release of less than 30% over 48 h. When exposed to a mildly acidic environment (pH 6.5), the cumulative release remained below 50%. However, under simulated TME conditions featuring pH 6.5 and 100 μM H2O2, the DOX release profile demonstrated significant time-dependent enhancement, plateauing after 24 h with a cumulative release reaching up to 80%. These results demonstrated that MnO2@CLDOX maintained excellent stability under normal physiological conditions, effectively preventing drug leakage and enabling more efficient tumor-targeted delivery. Upon reaching the tumor site, the MnO2 shell disintegrated in response to acidic pH and elevated H2O2 levels, accelerating DOX release. This dual-responsive behavior demonstrates the excellent TEM-responsiveness and efficient drug release performance of MnO2@CLDOX.
As mentioned before, MnO2-based NPs demonstrated the capacity to catalyze the decomposition of acidic H2O2, enabling efficient O2 generation (Equation 1). The O2-generating capacity of MnO2@CLDOX was also investigated in solutions. As illustrated in Figure 2B, we confirmed that MnO2@CLDOX could not produce O2 in pH 7.4 without adding H2O2. After introducing H2O2, abundant O2 was generated by MnO2@CLDOX, which proved the H2O2-triggered O2 generation ability of NPs. Compared with neutral conditions, an acidic environment (pH 6.5) enhanced O2-generating ability of MnO2@CLDOX. Collectively, these results indicate that MnO2@CLDOX can remain stable under physiological conditions and selectively responded to the environments with high level of H2O2 and H+ to produce O2 efficiently.
To investigate the charge reversal characteristics of MnO2@CLDOX, surface potential variations under different solution conditions were monitored using a zeta potential analyzer. As shown in Figure 2C, DOTAP (a cationic lipid) confers CLDOX with positive charges in acetate buffer solutions. However, after coating with MnO2, the nanomedicines (MnO2@CLDOX and MnO2@CL) exhibit negative surface charges, which reduces their uptake by normal tissues and is expected to enhance in vivo safety. Furthermore, when exposed to a TME-mimicking solution (pH 6.5, containing 100 μM H2O2), the nanomedicine exhibited TME-responsive charge reversal from negative to positive (Figure 2D). Generally, positively charged nanomedicines are more readily internalized by cells. This charge reversal capability facilitates higher chemotherapeutic drug delivery efficiency in tumor tissues, establishing a crucial pharmaceutical foundation for achieving low toxicity and high therapeutic efficacy in antitumor treatment.

Cellular uptake
To explore the in vitro cellular uptake profile of MnO2@CLDOX, MDA-MB-231 cells were employed. CLSM was utilized to detect the fluorescence of DOX, thereby tracing the drug molecules. As depicted in Figure 3, with the extension of incubation time, the red fluorescence in cell nuclei of the free DOX-treated group continuously increased. In contrast, relatively weaker red fluorescence was observed at all time points following MnO2@CLDOX treatment. This suggests that hydrophilic small-molecule drugs are more readily internalized by cells, potentially due to the multi-step processes required for nanodrugs, including endocytosis, disintegration, and gradual drug release, before nuclear accumulation. Notably, even after 12 h of incubation with MnO2@CLDOX, the intracellular drug signal remained relatively weak—likely due to the negative surface charge of the nanocarrier and incomplete disintegration of its MnO2 outer layer. In contrast, cells treated with CLDOX lacking MnO2 coating exhibited enhanced drug uptake. Further experiments under pH 6.5 and 100 μM H2O2 conditions demonstrated a significant increase in DOX uptake for MnO2@CLDOX. This revealed that the disintegration of the MnO2 outer layer under simulated TME conditions exposed CLDOX, thereby boosting cellular uptake via electrostatic interactions.
Furthermore, a semi-quantitative analysis of drug uptake across different nanoformulations was conducted using flow cytometry. As illustrated in Figures 4A and 4B, under simulated TME conditions characterized by acidity and oxidative stress, the MnO2@CLDOX group demonstrated significantly higher cellular drug accumulation at 12 h compared to that under physiological conditions, with uptake levels comparable to those of the CLDOX group. These findings demonstrated that MnO2@CLDOX could disintegrate and release CLDOX in a TME-responsive manner, achieving charge reversal to enhance tumor cell uptake of the drug.

Enhanced anticancer efficacy in vitro
The anticancer efficacy of MnO2@CLDOX in vitro was evaluated against MDA-MB-231 cells by MTT assay under simulated TME conditions. As shown in Figure 5A, the MnO2@CL showed no significant effect on cell viability at varying concentrations, indicating negligible antiproliferative activity of the blank carrier. In contrast, all other formulations demonstrated concentration-dependent cytotoxicity after 48-h treatment, with cell survival rates progressively decreasing as drug concentrations increased. Notably, at equivalent concentrations, the MnO2@CLDOX-treated group showed significantly lower cell viability than the CLDOX-treated group. These data demonstrated that the MnO2-based nanocarrier significantly potentiated the antiproliferative efficacy of DOX in tumor cells. To further validate the broad applicability of MnO2@CLDOX, its cytotoxicity was concurrently assessed against 4T1 and MCF-7 breast cancer cells under simulated TME conditions. Similar to the results observed in MDA-MB-231 cells, the MnO2@CLDOX-treated groups exhibited markedly lower cell viabilities compared to the CLDOX-treated groups across these cell lines (Figure S5). According to Figure 4B, CLDOX and MnO2@CLDOX showed no significant difference in cellular uptake, and MnO2@CL alone did not exhibit notable cytotoxicity toward cells. We therefore speculate that hydroxyl radicals generated via MnO2-mediated Fenton-like reactions may exert a sensitizing effect on the chemotherapeutic efficacy of DOX.26
To further explore the capacity of MnO2@CLDOX to induce apoptosis under simulated TME conditions, the Annexin V-APC/7-AAD staining was used. As revealed by Figures 5B and 5C, the total percentage of apoptotic cells (including early and late apoptotic cells) induced by MnO2@CLDOX reached 48% in MDA-MB-231 cells, far exceeding the findings in CLDOX and MnO2@CL-treated groups, which demonstrated that the introduction of MnO2 could bolster chemotherapy-induced cell apoptosis.
By establishing a 3D MCTS model of MDA-MB-231 cells, we assessed the growth inhibitory effects of nanodrugs on solid tumors.27 As illustrated in Figures 5D and 5E, after 9 days of drug treatment on MCTSs (with identical initial volumes), both the control group and the MnO2@CL group exhibited rapid MCTS volume expansion, demonstrating a lack of tumor cell proliferation inhibitory activity. In contrast, the remaining three treatment groups showed slower growth rates and ultimately smaller final volumes. Compared with the DOX and CLDOX-treated groups, the MnO2@CLDOX-treated group displayed the most pronounced inhibitory effect on MCTS growth post-incubation. These results confirm that the MnO2 and DOX components in the liposomal nanosystem synergistically inhibit tumor growth in the 3D MCTS level.

In vitro mechanism studies
The influence of MnO2@CLDOX on the intracellular O2 was assessed using a commercial hypoxia probe, [Ru(dpp)3]Cl2.28 Under hypoxic conditions, untreated MDA-MB-231 cells exhibited the strongest red fluorescence, indicating the lowest O2 availability. Following 2-h incubation with MnO2@CL or MnO2@CLDOX under pH 6.5 and 100 μM H2O2 conditions, intracellular O2 level increased significantly; however, no O2 production was observed in CLDOX-treated cells (Figures 6A and 6B). This phenomenon can be attributed to the robust catalytic activity of MnO2 in O2 generation. To further investigate the oxygen-regulated expression HIF-1α protein following different treatments under simulated TME conditions, western blot analysis was performed. As shown in Figure 6C, HIF-1α expression remained relatively low under normoxic conditions. In hypoxic environments, both the control group and the CLDOX group exhibited elevated HIF-1α protein levels. Conversely, the MnO2@CL and MnO2@CLDOX groups demonstrated significantly reduced HIF-1α expression, indicating enhanced intracellular O2 levels. These findings confirm that MnO2@CLDOX can effectively elevate tumor cell oxygenation to alleviate hypoxic TME.
It is well established that tumor hypoxia modulates the antitumor efficacy of chemotherapy by regulating hypoxia-inducible factor-1 (HIF-1) pathways.29 CTGF and collagen I, which are regulated by HIF-1, function as key regulators of collagen deposition in tumor tissue—this process can hinder drug penetration. Based on the hypoxia relief capacity of MnO2@CLDOX, we speculated that MnO2@CLDOX could downregulate the expression of CTGF and collagen I via O2 production. As illustrated in Figure 6C, under normoxic conditions, both CTFG and collagen Ⅰ expression levels were low, indicating that oxygen suppresses the HIF-1/CTFG/collagen Ⅰ signaling pathway. Under hypoxic conditions, the control group and CLDOX group exhibited elevated expressions of CTFG and collagen Ⅰ proteins. Conversely, MnO2@CL and MnO2@CLDOX treatment groups demonstrated suppressed expression of these proteins, suggesting that increased intracellular O2 levels effectively downregulate HIF-1α downstream protein expression. These findings confirm that the oxygen generated by MnO2@CLDOX alleviates tumor hypoxia, thereby inhibiting the HIF-1/CTFG/collagen Ⅰ pathway associated with drug penetration barriers. This mechanism explains how the nanocarrier enhances chemotherapeutic drug delivery efficiency and potentiates antitumor efficacy.
The tumor tissue penetration capability of MnO2@CLDOX was evaluated by detecting DOX red fluorescence in MCTS at different depths using CLSM. As shown in Figure 6D, hydrophilic small-molecule DOX demonstrated strong penetration capacity within 24 h, exhibiting intense fluorescence signals across all depth sections of the MCTS. Compared with DOX, CLDOX could not penetrate completely and uniformly into the deep layers of MCTS. The likely reason is that the positively charged CLDOX may induce interactions with negatively charged components on the surface of MCTS, thereby hindering its penetration depth and uniformity. In contrast, although the MnO2@CLDOX could penetrate well into the interior of MCTS, the fluorescence intensity is relatively weak. This may be due to the surface negative charge and the insufficient degradation of MnO2 within a short period. To verify this hypothesis, MCTSs were further cultured with MnO2@CLDOX under conditions of pH 6.5 and 100 μM H2O2. Remarkably, enhanced drug signals were observed throughout the depth sections after 24-h incubation, indicating that the nanoformulation can respond to TME to enhance DOX penetration. This phenomenon is presumably due to: (1) MnO2 component disintegration facilitating small-molecule DOX release and (2) MnO2-mediated oxygen generation through reaction with H2O2, which may reduce stromal collagen synthesis to improve drug diffusion.

In vivo biodistribution and pharmacokinetics study
The MDA-MB-231 and 4T1 cells xenograft model was used to assess the In vivo distribution and pharmacokinetics study of MnO2@CLDOX. Initially, the near-infrared fluorescent probe DiR was encapsulated into the nanoformulations to track their real-time distribution in vivo.30 As displayed in Figure 7A, free DiR undergoes rapid metabolic clearance, with its fluorescence intensity gradually diminishing over time following tail vein injection. The fluorescence of free DiR is predominantly distributed in the hepatic region, and ex vivo tissue fluorescence analysis post 48 h reveals negligible fluorescent signals within tumor tissues. In contrast, MnO2@CLDOX/DiR demonstrated significantly more vigorous fluorescence intensity than free DiR at the same time post-injection, confirming their prolonged systemic circulation properties. Moreover, the fluorescence intensity of MnO2@CLDOX/DiR at the tumor site increased progressively over time, with a strong fluorescent signal still detectable at 48 h post-injection, confirming its excellent passive tumor-targeting capability. Similarly, the Mn biodistribution assay (Figure 7B) demonstrates that MnO2@CLDOX exhibits predominant accumulation in tumor tissue—with peak accumulation occurring at 12 h post-injection—alongside the liver and spleen, while showing minimal accumulation in other normal organs (hearts, lungs, kidneys). The prolonged tumor accumulation of MnO2@CLDOX can be attributed to the dual mechanisms of EPR effect and MnO2-mediated TME modulation. Interestingly, due to its positive charge, CLDOX/DiR exhibits a significantly faster in vivo clearance rate than MnO2@CLDOX/DiR, with predominant accumulation in the hepatic region and weak fluorescent signals in tumor sites. The fluorescence of ex vivo tissue post 48 h reveals CLDOX/DiR shows minimal accumulation in tumors, with most of the distribution observed in the liver, spleen, lung, and kidney.31
Figure 7C reveals that MnO2@CLDOX possessed more favorable pharmacokinetic profiles than free DOX and CLDOX, manifesting extended circulation duration and diminished systemic clearance. These results indicated that coating CLDOX with MnO2 significantly prolongs its systemic circulation, enhances tumor-targeting efficacy, reduces drug accumulation in normal tissues, and mitigates toxic effects on healthy tissues.

Enhanced anticancer efficacy in vivo
To further evaluate the in vivo therapeutic efficacy of MnO2@CLDOX, treatments were initiated once the average tumor volume reached approximately 120 mm3. As illustrated in Figures 8A–8C, the tumor growth rates in the saline group, MnO2@CL group, and CLDOX group exhibited more rapid increases. In contrast, the growth rate in the MnO2@CLDOX group started to decelerate gradually. Notably, the tumor weight in the MnO2@CLDOX-treated group was significantly lower than that in the other groups (Figure 8D). According to Figure 8E, the tumor inhibition rate of MnO2@CLDOX was approximately 73.1%, which was significantly higher than those of MnO2@CL (10.2%) and CLDOX (43.8%). Collectively, these results demonstrate that the TME-responsive charge-reversible nanocarrier enhances the delivery efficiency of chemotherapeutic drugs and potentiates in vivo antitumor efficacy.
Immunohistochemical analysis of tumor sections was further performed to assess the expression of HIF-1α, collagen I, and CTGF proteins. Consistent with the in vitro findings, the HIF-1α signal was markedly downregulated in the oxygen-generating groups (Figure 9A)—this confirms the in vivo hypoxia-alleviating effect of MnO2@CL and MnO2@CLDOX. Moreover, as shown in Figures 9B and 9C, the protein expression levels of collagen I and CTGF were significantly lower in the MnO2@CL and MnO2@CLDOX groups than in the other two groups. Semi-quantitative statistical analysis of the positive staining area and optical density revealed that the differences between MnO2@CL and MnO2@CLDOX groups reached statistical significance, which provided reliable quantitative evidence for the results of qualitative observation (Figure S7). This reduction facilitates the deep penetration of the drug into the tumor interior. As shown by the TUNEL staining assay (Figure 10), marked apoptosis was observed in tumor tissues following treatment with MnO2@CLDOX. In contrast, other therapeutic groups (CLDOX and MnO2@CL) did not exhibit such a pronounced effect.
2To evaluate the potential toxicity of MnO2@CLDOX treatment, body weights of mice from different groups were monitored within 13 days. No noticeable weight loss was detected in all treated mice (Figure S6). Serum biochemical tests were performed further to evaluate the renal and hepatic toxicity of MnO2@CLDOX. As shown in Figures S8A–S8D, there were no significant changes in the markers associated with liver function (ALT and AST), kidney function (BUN and CREA). Moreover, no obvious toxicity induced by MnO2@CLDOX was observed via blood routine test indicators, as shown in Figure S9. Finally, histological analysis of major normal organs (heart, liver, spleen, lungs, and kidneys) confirmed high biosafety of MnO2@CLDOX, with minimal tissue damage observed in treated mice (Figure S10). Taken together, these results verified that this MnO2-coated nanoliposome exhibited efficient in vivo therapeutic performance with negligible systemic toxicity.

Discussion
In summary, we constructed a TME-responsive charge-reversible nanocarrier to enhance DOX chemotherapy by optimizing drug transport to cancer cells through multiple mechanisms. Innovatively, the in situ reaction between MnCl2 and CaO2 NPs was harnessed to coat cationic liposomes loaded with DOX (CLDOX) with a negatively charged MnO2 layer, forming the MnO2@CLDOX nanosystem. Following intravenous administration, this nanoformulation exhibited prolonged blood circulation and reduced accumulation in normal tissues in vivo. Upon reaching tumors via the EPR effect, the outer MnO2 layer undergoes acid/H2O2-responsive degradation, releasing oxygen and CLDOX. The positively charged CLDOX facilitates enhanced cellular uptake by tumor cells, while the released oxygen downregulates the HIF-1α/collagen I/CTGF pathway, thereby reducing ECM deposition and promoting the deep penetration of drugs within tumor tissues. Importantly, the MnO2-coated cationic liposomal carrier not only serves as a versatile platform for delivering various chemotherapeutics or small interfering RNAs, but ‌also, by virtue of its Mn2+-releasing property, is inherently endowed with theranostic potential for possible imaging-guided therapy. Overall, this MnO2-coated cationic liposome emerges as a promising theranostic strategy with broad applicability in precision cancer therapy, integrating efficient drug delivery, TME modulation, and noninvasive imaging for translational applications.

Limitations of the study
Despite the promising antitumor potential of the MnO2@CLDOX nanosystem, several limitations need to be addressed: the degradation rate of MnO2 and the kinetics of O2 release in the TME remain uncontrollable, weakening the precision of hypoxia modulation. Additionally, the study does not investigate the MRI imaging potential enabled by Mn2+ release, overlooking the theranostic value of the nanosystem. Lastly, the absence of active targeting ligands on the nanocarrier surface results in insufficient specific internalization by tumor cells and increases the risk of off-target effects.

Resource availability

Lead contact
Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Xiaojuan Zhang (xjzhang@zjxu.edu.cn).

Materials availability
This study did not generate new unique reagents.

Data and code availability
This article does not report original code. All data associated with this study are present in the article or the supplemental information. All data reported in this article will be shared by the lead contact upon request. Any additional information required to reanalyze the data reported in this article is available from the lead contact upon request.

Acknowledgments
This work was supported by 10.13039/501100004731Zhejiang Provincial Natural Science Foundation (nos. ZCLQ24H3001, LQN25H160047), Zhejiang Provincial Medical and Health Science and Technology Program (no. 2024KY1692), the Science and Technology Bureau of Jiaxing (2024AY40030, 2024AY40013, 2024AY10009, and 2024AY10042) and the “Innovative Jiaxing · Excellent Talent Support Program”-Top Talents in Technological Innovation.

Author contributions
C.H. and X.Z.: writing – review and editing, writing – original draft, investigation, and conceptualization; S.K.: writing – original draft, methodology, and investigation; J.Z.: methodology; L.C. and G.Z.: conceptualization and methodology; L.J. and B.D.: conceptualization, methodology.

Declaration of interests
The authors declare no competing interests.

STAR★Methods

Key resources table

Experimental model and study participant details

Animals
Female BALB/c nude mice (4-5 weeks old) were purchased from Beijing Huafukang Bioscience Technology Co., Ltd. (Beijing, China). The animals were housed in sterile filter-top cages under specific pathogen-free (SPF) conditions, provided with autoclaved standard rodent chow and fresh water ad libitum, and maintained on a 12-h light/dark cycle. This breast cancer study exclusively employed female experimental models, with sex as a consistent controlled variable. No gender-related factors were included in the study, and therefore sex/gender had no impact on the observed experimental results. All experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Jiaxing University (SYXK2020-0007).

Cell lines
MDA-MB-231, MCF-7 (human breast cancer cells) and 4T1 cells (mouse breast cancer cells) were obtained from Wuhan Sunncell Biotechnology Life Co., Ltd. These cell lines were authenticated by the supplier via STR profiling (human cells) and species-specific identification (murine cells), with the authentication certificate provided. All cell lines were tested for mycoplasma contamination by the supplier (confirmed negative). The cells were cultured in basal medium, supplemented with 10%FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. The hypoxic cell incubation system was purchased from PUHE Biotechnology Co., Ltd. (Wuxi, China), capable of generating a humidified hypoxic atmosphere maintained at 37°C with 5% CO2 and 1% O2.

Method details

Preparation of MnO2@CLDOX
A 10-mL microemulsion was prepared by combining cyclohexane/Igepal CO-520 (71:29, v/v) containing 500 μL of 25% ammonium hydroxide (NH3·H2O) and 60 μL of DSPA, followed by vigorous magnetic stirring at room temperature. Concurrently, a second 10-mL microemulsion was generated by homogenizing cyclohexane/Igepal CO-520 (71:29, v/v) with 250 μL of 30% H2O2, 250 μL of 4 M CaCl2 solution, and 5 mg of DOX under similar stirring conditions. Both microemulsions were magnetically stirred for 10 min. Subsequently, the two microemulsions were combined and stirred for 30 min to complete the reaction. To obtain DSPA@DOX-CaO2, 15 mL of ethanol was employed to disrupt the microemulsion and precipitate the NPs, which were subsequently collected, washed, and redispersed in 15 mL of chloroform. Afterwards, 300 μL of DOTAP, 100 μL of CHOL and 50 μL of DSPE-PEG in chloroform were added to 3 mL of DSPA@DOX-CaO2 solution. After rotary evaporation, a thin film was formed, adding 3 mL of MnCl2 solution (1 M) and sonification. After reaction for 5 min, the final brownish product MnO2@CLDOX was collected by centrifugation.

Characterization of MnO2@CLDOX
The morphology was characterized using by transmission electron microscopy (TEM). The UV-vis absorption spectra of the NP solutions were recorded using a spectrophotometer in a quartz cuvette over the wavelength range of 300-800 nm at room temperature. The hydrodynamic diameter and zeta potential of the NPs were determined by dynamic light scattering (DLS) measurements after the sample were diluted in deionized water. The DOX loading content in MnO2@CLDOX was quantified by high-performance liquid chromatography (HPLC) following dissolution of the samples in an appropriate solvent and filtration. The Mn content was analyzed by inductively coupled plasma atomic absorption spectrometry (ICP-AAS) after acid digestion of the samples, using a standard calibration curve method. The dissolved O2 concentration was measured by a JPSJ-605F dissolved oxygen meter at 37°C.

Cellular uptake
The red fluorescence of DOX was utilized to track the intracellular localization of NPs in cells. Briefly, cells were plated into 12-well plates at a seeding density of 2×105 cells per well. Following 24 h of culture, the cells were treated with free DOX, CLDOX, and MnO2@CLDOX formulations, each containing DOX at a final concentration of 1 μM. After incubation for specified time intervals, the culture medium was removed, and the cell monolayers were washed three times with phosphate-buffered saline (PBS, pH 7.4) to remove unbound materials. Subsequently, the cells were counterstained with 4′,6-diamidino-2-phenylindole (DAPI, 1.2 μg/mL in PBS) for 5 min to visualize the nuclei, followed by gentle rinsing with PBS to remove excess stain. Fluorescence imaging was performed using confocal laser scanning microscopy (CLSM), with appropriate excitation/emission settings for DOX (λex = 488 nm, λem = 588 nm) and DAPI (λex = 405 nm, λem = 461 nm).

Cytotoxic activities
Cell viability following different treatments was evaluated by measuring mitochondrial dehydrogenase activity via the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Briefly, cells were plated into 96-well plates at a seeding density of 4×103 cells per well and incubated overnight to allow attachment. The cells were then treated with various concentrations of test formulations (MnO2@CL, CLDOX, MnO2@CLDOX) for 48 h at pH 6.5 and under 100 μM H2O2 conditions. Following treatment, the culture medium was carefully removed, and 20 μL of MTT solution (5 mg/mL in PBS, filter-sterilized) was added to each well. After a 4-h incubation at 37°C, the supernatant was discarded, and 150 μL of dimethyl sulfoxide (DMSO) was added to solubilize the formazan crystals. The absorbance of the resulting solution, reflecting viable cell number, was measured at 570 nm with a reference wavelength of 630 nm using a microplate reader (VersaMax, USA). All experimental conditions were performed in quintuplicate, and data are presented as mean ± standard deviation (SD) from three independent experiments.
For apoptosis analysis, cells were seeded on 6-well plates and cultured in DMEM with 10% FBS for 12 h. Then the medium was replaced with fresh medium and different drug formulations were added (DOX concentration: 1 μM) for 24-h incubation at pH 6.5 and under 100 μM H2O2 conditions. The cells were collected and stained with an Annexin V- APC/7-AAD Apoptosis Detection Kit (KeyGen Biotech). A BD Accuri C6 Flow Cytometer was used for analysis.
The formation of multicellular tumor spheroids (MCTS) was monitored using an optical microscope (TE2000-S, Nikon, Japan). For MCTS inhibition assays, tumor spheroids with a diameter of approximately 200 μm were grouped (n=5 per group). Selected spheroids were treated with different formulations and incubated at 37°C for 9 days. Their diameters were imaged and recorded every 2 days via an optical microscope.

Drug penetration in MCTS
Tumor spheroids with diameters of approximately 400 μm were treated with free DOX, CLDOX, MnO2@CLDOX, and MnO2@CLDOX + H2O2 + H+ (each at a DOX concentration of 2 μM) for 24 h, respectively. Following incubation, the tumor spheroids were washed three times with ice-cold PBS, fixed with paraformaldehyde for 30 min, and mounted on cavity microscope slides. CLSM was used to acquire Z-stack images of the tumor spheroids, with optical sections collected at 20 μm intervals from the surface of the spheroid to its midsection.

Intracellular O2 determination
Intracellular oxygen levels were assessed using the ruthenium-based optical probe [Ru(dpp)3]Cl2. Briefly, MDA-MB-231 cells were first incubated under hypoxic conditions (1% O2) for 24 h, followed by the addition of [Ru(dpp)3]Cl2 (10 μg/mL) and further incubation for an additional 12 h. The cells were then subjected to treatments for 2 h under simulated TME conditions (pH 6.5 and 100 μM H2O2). After treatment, the cells were washed twice with PBS, and intracellular O2 levels were evaluated using fluorescence microscopy by detecting the fluorescence of [Ru(dpp)3]Cl2 (λex = 450 nm, λem = 610 nm).

Western blot analysis
Total cellular proteins were extracted with RIPA buffer. Protein samples (30 μg) were separated via 10% SDS-PAGE and transferred onto PVDF membranes. Following blocking with 5% skimmed milk for 1 hour, the membranes were incubated with primary antibodies overnight at 4°C, rinsed with TBST, and then probed with HRP-conjugated secondary antibodies for 1 h at room temperature. Following additional washes, signals were detected using an enhanced chemiluminescence system. β-Tubulin was used as a loading control. Antibodies used for western blotting included those against HIF-1α (1:1000, Cell Signaling), CTGF (1:1000, Cell Signaling), and Collagen I (1:1000, Cell Signaling).

In vivo experiments
In vivo imaging was performed on MDA-MB-231 xenograft tumor-bearing mice using a noninvasive optical imaging system. The hydrophobic fluorescent dye DiR was loaded into MnO2@CLDOX or CLDOX via the film dispersion. Female BALB/c nude mice were intravenously administered MnO2@CLDOX/DiR, CLDOX@DiR, or free DiR at 5 μg DiR per mouse. Fluorescence intensity images of each mouse were acquired at 0.5, 2, 6, 12, 24, and 48 h post-injection.
For tissue Mn concentration determination, 4T1 tumor-bearing xenograft mice received a single intravenous dose of MnO2@CLDOX (5 mg/kg DOX equivalent) via intravenous injection, followed by sacrifice at 1, 4, 12 and 24 h post-administration (n = 4 per group). The heart, liver, spleen, lung, kidney and tumor tissues were collected, digested with concentrated nitric acid, diluted to 2 mL with Milli-Q water, and subjected to ICP-AAS for Mn quantification.
To evaluate the pharmacokinetic profiles, free DOX, CLDOX, and MnO2@CLDOX (equivalent to 5 mg/kg DOX) were intravenously administered via the tail vein to female BALB/c mice (n = 4 per group). At 0.5, 1, 3, 6, 12, 24, and 48 h post-injection, 10-20 μL of blood was collected from the tail vein and immediately mixed with heparin sodium solution (1000 U/mL) to prevent coagulation, followed by dilution with PBS. The blood samples were centrifuged to obtain plasma, and the DOX concentration in plasma was quantified by HPLC with fluorescence detection for subsequent pharmacokinetic analysis.
To evaluate the in vivo antitumor efficacy of MnO2@CLDOX, MDA-MB-231 xenograft tumor-bearing mice were randomly assigned to 4 groups (n = 5 per group). When the average tumor volume reached approximately 120 mm3, the mice received tail vein injections of saline, CLDOX, MnO2@CL, and MnO2@CLDOX at a dose of 5 mg/kg DOX or the equivalent dose for formulations without DOX. Following treatment, tumor volumes and mouse body weights were measured every two days. On day 13 post-treatment, the mice were euthanized, and major organs as well as tumor tissues were collected for subsequent analysis.

Quantification and statistical analysis

Immunohistochemistry quantification
Immunohistochemical staining of HIF-1α, collagen I and CTGF was quantitatively evaluated in a double-blinded fashion. Three random microscopic fields were captured per section. The staining intensity was scored as 0 (negative), 1 (weak), 2 (moderate), and 3 (strong). The H-score was calculated as ∑(intensity score × percentage of positive cells), and ImageJ software was used for auxiliary quantitative analysis.

Statistical analysis
All statistical analyses were performed using GraphPad Prism (version 10.2.0) unless otherwise specified. Two-sided non-parametric t-test (Mann-Whitney) or unpaired t-test were used to assess differences between conditions. When more than two groups were compared, we used non-parametric one-way ANOVA (Kruskall-Wallis). For the assessment of the effect of two factors (for the spatial analysis) we used a two-way ANOVA. Statistical tests used, n numbers and p values are displayed in the appropriate figures and figure legends. P values < 0.05 were considered statistically significant, with asterisks denoting the level of significance: ∗P ≤ 0.05, ∗∗P ≤ 0.01, ∗∗∗P ≤ 0.001, ns means not significant.