Dual-Sensitive Charge-Conversional Polymeric Prodrug for Efficient

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Dual-sensitive charge-conversional polymeric prodrug for efficient co-delivery of demethylcantharidin and doxorubicin Yanjuan Wu, Dongfang Zhou, Qingfei Zhang, Zhigang Xie, Xuesi Chen, Xiabin Jing, and Yubin Huang Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.6b00705 • Publication Date (Web): 06 Jul 2016 Downloaded from http://pubs.acs.org on July 7, 2016

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Dual-sensitive charge-conversional polymeric prodrug for efficient co-delivery of demethylcantharidin and doxorubicin Yanjuan Wua,b, Dongfang Zhou*a, Qingfei Zhanga,b, Zhigang Xiea, Xuesi Chenc, Xiabin Jinga, Yubin Huang*a a

State Key Laboratory of Polymer Physics and Chemistry, Changchun Institute of Applied

Chemistry, Chinese Academy of Sciences, Changchun 130022, People’s Republic of China b

University of Chinese Academy of Sciences, Beijing 100049, People’s Republic of China

c

State Key Laboratory of Polymer Ecomaterials, Changchun Institute of Applied Chemistry,

Chinese Academy of Sciences, Changchun 130022, People’s Republic of China KEYWORDS: Dual-sensitive; charge-conversion; polymeric prodrug; co-delivery; drug resistance; nanoparticles. ABSTRACT: Tumor is a complicated system, and tumor cells are typically heterogeneous in many aspects. Polymeric drug delivery nano-carriers sensitive to a single type of bio-signals may not release cargos effectively in all tumor cells, leading to low therapeutic efficacy. To address the challenges, here, we demonstrated a pH/reduction dual-sensitive charge-conversional polymeric prodrug strategy for efficient co-delivery. Reduction-sensitive disulfide group and acid-labile anticancer drug (demethylcantharidin, DMC)-conjugated β-carboxylic amide group were repeatedly and regularly introduced into copolymer chain simultaneously via facile CuAAC click

polymerization.

The

obtained

multifunctional

polymeric

prodrug

P(DMC),

mPEG-b-poly(disulfide-alt-demethylcantharidin)-b-mPEG was further utilized for DOX

ACS Paragon Plus Environment

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encapsulation. Under tumor tissue/cell microenvironments (pH 6.5 and 10 mM GSH), the DOX-loaded polymeric prodrug nanoparticles (P(DMC)@DOX NPs) performed surface negative-to-positive charge conversion and accelerated/sufficient release of DMC and DOX. The remarkably enhanced cellular internalization and cytotoxicity in vitro, especially against DOX-resistant SMMC-7721 cells were demonstrated. P(DMC)@DOX NPs in vivo also exhibited higher tumor accumulation and improved antitumor efficiency compared to P(SA)@DOX NPs with one drug and without charge-conversion ability. The desired multifunctional polymeric prodrug strategy brings a new opportunity for cancer chemotherapy. 1. INTRODUCTION Over the past decades, polymeric drug delivery nano-carriers have attracted increasing attention due to the improved pharmacokinetics and higher tumor accumulation profile via the enhanced permeation and retention (EPR) effect.1-6 To further improve the specificity to disease sites, great efforts have been paid to develop controlled drug release systems in response to tumor tissue/cell microenvironments.7-10 Typical biological stimuli exploited include pH, reductive potential, enzymes, oxidative stress and temperature.11-14 Among these, pH and redox-sensitivity are often employed to trigger drug release from polymeric nanoparticles.15,16 Compared to normal physiological conditions (pH ~ 7.4, GSH ~ 2.0 to 20 µM), tumor tissue/cell usually contains more acidic extracellular environment (pH ~ 6.5 to 7.0) and higher GSH level (~ 0.5 to 10 mM).17-19 As disulfide possesses reduction-sensitive property, multifarious disulfide containing polymeric nano-carriers have been extensively studied.20,21 Particularly, disulfide


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arranged regularly in polymer backbone can induce rapid and complete degradation.22,23 pH-sensitive polymeric nano-carriers usually incorporate orthoester, acetal and hydrazone groups as acid-labile linker, which can release cargos in the endosomal/lysosomal compartments.8,17 Recently, several reduction and pH dual-responsive polymeric drug/gene delivery systems have been reported.24-28 For instance, Zhong’s group developed a pH and redox dual-sensitive biodegradable polycarbonate micelles for efficient DOX encapsulation and triggered intracellular release.24 Shuai and co-workers synthesized a triblock copolymer for co-delivery of DOX and siRNA.26 As a further evolution, charge-conversional polymeric nano-carriers responding to tumor microenvironment acidity have been applied recently not only to prolong blood circulation time but also to enhance cellular uptake.29-31 Typically, β-carboxylic amides have shown to be effective to shield positively charged surfaces during blood circulation to minimize nonspecific adsorption, whereas positive charge recovers from negative under acidic pH nearby tumor site to facilitate cancer cell internalization and endosome/lysosome escape.32-34 Although promising, challenges still exist. Polymeric nano-carriers are commonly designed to respond to only one type of tumor tissue/cell bio-signals to trigger cargo release, mostly the intracellular GSH, lysosomal acidity, or reactive oxygen species (ROS). As we all known, tumor is a complicated system, and tumor cells are typically heterogeneous in many aspects.35 The above bio-signals may present in different tumors, but may also exist in the distinct cells of one tumor, and even in one cancer cell at different stages. Therefore, polymeric nano-carriers responsive to a single type of bio-signals may not release cargo effectively in all the fraction of tumor cells, leading to low therapeutic efficacy. Multi-sensitive polymeric nano-carriers, which


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could realize controllable drug release and obtain excellent therapeutic outcome, have received extensive attention.16 However, elaborate design and synthesis, careful protection and de-protection, or post-polymerization modifications are usually necessary.36 The development of multi-functional “smart” polymeric nanoparticles via ordinary design and simple synthesis is the still the remained immense challenge. Taking advantage of the discrepancy between healthy tissue and tumor tissue/cell microenvironments,

herein,

we

reported

a

novel

dual-sensitive

(pH/reduction)

charge-conversional polymeric prodrug strategy for efficient co-delivery. Disulfide-incorporated dialkyne monomer and β-carboxylic amide/anticancer drug (demethylcantharidin, DMC)-bearing diazide monomer were designed for facile CuAAC click polymerization. DMC is an inhibitor of serine/threonine protein phosphatase 2A (PP2A), a promising therapeutic target for cancer treatment.37

The

obtained

ABA-type

triblock

multifunctional

polymeric

prodrug

mPEG-b-poly(disulfide-alt-demethylcantharidin)-b-mPEG, P(DMC), was further utilized for DOX encapsulation (Scheme 1). The drug-loaded multifunctional polymeric prodrug nanoparticles (P(DMC)@DOX NPs) have several advantages: (1) repeated acid-labile anticancer drug (DMC)-conjugated β-carboxylic amide group on the polymer sidechains could realize tumor extracellular pH-induced DMC release and negative-to-positive charge conversion for better tumor accumulation/cellular internalization simultaneously; (2) disulfide bonds arranged regularly in the backbone of polymer would also induce complete intracellular dissociation of NPs for controlled and sufficient DOX release; (3) conjugated DMC and encapsulated DOX for efficient combination chemotherapy; (4) multifunction was involved in one polymeric


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nano-carrier system by elaborated design and facile click chemistry; (5) drug-conjugated monomer for polymerization to obtain high drug loading content (DLC) and drug loading efficiency (DLE). Another similar structured copolymer, P(SA) with succinic anhydride instead of DMC, was also synthesized as a control.

Scheme 1 Schematic illustration of pH/reduction dual-sensitive polymeric prodrug nanoparticles with tumor extracellular pH triggered charge-conversion ability for efficient intracellular co-delivery of DMC and DOX. 2. EXPERIMENTAL SECTION


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2.1. Materials 2.2’-Dithiobis[1-(prop-2-ynylcarbamoyl)-ethyl-carbamicacidtert-butyl

ester]

(dialkyne

monomer) and azide-terminated mPEG5k were synthesized according to our previous work.38 Demethylcantharidin (DMC), succinic anhydride, glutathione, and buthionine-sulfoximine (BSO) were purchased from Aladdin Chemistry Co. Ltd (Shanghai, China). Poly(ethylene glycol) methyl ether with an Mn of ca. 5000, 3-(4,5-Dimethyl-thiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) and Hoechst 33258 were obtained from Sigma-Aldrich. Cuprous chloride (CuCl), N,N,N′,N″,N″-pentamethyldiethylenetriamine (PMDETA) were purchased from Energy Chemical Co. Ltd (Shanghai, China). Doxorubicin hydrochloride (DOX·HCl) was obtained from Zhejiang Hisun Pharmaceutical Co. Ltd. Dimethylformamide (DMF) and triethylamine (TEA) were maintained with calcium hydride (CaH2) and purified by distillation. 2.2. Measurements Proton and carbon nuclear magnetic resonance (1H and 13C NMR) spectra were collected on a Bruker AVANCE DRX 400 spectrometer in deuterated chloroform (CDCl3) or dimethyl sulfoxide (DMSO-d6). Fourier-transform infrared (FT-IR) measurements were performed on a Bruker Vertex70 Win-IR instrument. Electrospray ionization mass spectrometry (ESI-MS) measurement was conducted on a Watera Quattro Premier XE system. The molecular weight and polydispersity index of the synthetic copolymers were determined by a Waters 515 gel permeation chromatograph (GPC) instrument equipped with two linear Tskgel Super columns (AW5000 and AW3000), Waters 515 HPLC pump and an OPTILAB DSP Interferometric Refractometer detector (injection volume: 100.0 µL; column temperature: 50 °C; eluant: DMF


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containing 0.01 M LiBr; flow rate: 1.0 mL min-1; standard: polystyrene). Zeta potential measurements were determined on a Malvern Zetasizer Nano ZS. DLS measurements were performed on Brookhaven 90Plus size analyzer. TEM images were determined on a JEOL JEM-1011 transmission electron microscope with an acceleration voltage of 100 kV. The TEM samples were obtained by dropping 10 µL of solution on the copper grid coated with carbon and then drying in the air. DOX concentrations were measured by an ultraviolet-visible (UV-vis) spectrometer (UV-2450PC, Shimadzu, Kyoto, Japan). 2.3. Synthesis of copolymers 2.3.1. Synthesis of DMC-conjugated diazide monomer 2-azido-1-azidomethyl ethylamine was first synthesized following a previously published literature.39 Then, 2-azido-1-azidomethyl ethylamine (1.381 g, 9.34 mmol) and DMC (1.427 g, 8.49 mmol) were added to a flame dried 100 mL flask. Under argon, dried DMF (20 mL) and TEA (3 mL) were syringed. The mixture was stirred at 60 °C for 24 h. Then the mixture was concentrated under vacuum, and the resulting crude product was recrystallized from acetone to afford DMC-conjugated diazide monomer. Yield: 84%. 1H NMR (CDCl3, 400 MHz): δ (ppm) = 4.91 (m, 2H), 4.30 (m, 1H), 3.79 (q, 2H), 3.57 (q, 2H), 2.92 (d, 2H), 1.87 (m, 2H), 1.63 (m, 2H). 2.3.2. Synthesis of SA-conjugated diazide monomer Succinic anhydride (1.000 g, 10 mmol) and 2-azido-1-azidomethyl ethylamine (1.551 g, 11 mmol) were dissolved in 20 mL of dried DMF under argon with strong stirring, and then 2.8 mL of dried triethylamine (2-fold molar relative to that of succinic anhydride) was added. After reaction at 60 °C for 24 h, the reactive solution wasconcentrated in vacuum, and the resulting


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mixture was re-dissolved in 200 mL of ethyl acetate, washed for three times with saturated KHSO4 and NaCl. After drying the solution with anhydrous MgSO4 and evaporating solvent, SA-conjugated diazide monomer was obtained with 74% yield. 1H NMR (DMSO-d6, 400 MHz): δ (ppm) = 4.02 (m, 1H), 3.37 (d, 4H), 8.18 (d, 1H), 2.44 (t, 2H), 2.36 (t, 2H). 2.3.3. Synthesis of triblock copolymers P(DMC) and P(SA) CuAAC click polymerization proceeded under argon in Schlenk tube. CuCl was used as catalyst and PMDETA was applied as chelating agent. In general, dialkyne monomer (0.565 g, 1.1 mmol), DMC-conjugated diazide monomer (0.309 g, 1mmol) and PMDETA (0.070 g, 0.2mmol) were charged into a Schlenk tube, to which 10 mL of dried DMF was added. The solution was deoxidized by two freeze–pump–thaw cycles, and CuCl (0.021 g, 0.2mmol) was added to it. After two more freeze–pump–thaw cycles, the polymerization was carried out at 50 °C for 24 h. To ensure dialkyne end groups on the hydrophobic block, solution of dialkyne monomer (0.051 g, 0.1mmol) was further added to the tube at the end of polymerization. After precipitation in cold diethyl ether and filtration, the hydrophobic block was dialyzed against EDTA-2Na solution and pure water using a Spectra/Por Regenerated Cellulose membrane (MWCO 14000). Then, the obtained hydrophobic block (0.400 g), superfluous azide-terminated mPEG5k (0.801 g), PMDETA (0.055 g, 0.320 mmol) and CuCl (0.016 g, 0.160 mmol) were reacted in 20 mL of DMF at 50 °C for 48 h after a de-oxidation process. After dialyzed (MWCO 7000) against DMF, EDTA-2Na solution and pure water, the final polymer P(DMC) was obtained. The control copolymer mPEG5k-b-poly(disulfide-alt-succinic anhydride)-b-mPEG5k (P(SA)) was prepared in the same way, while SA-conjugated diazide monomer was used to


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instead of DMC-conjugated diazide monomer. 2.4. Preparation of blank NPs and DOX-loaded NPs Blank NPs were prepared using a modified nano-precipitation method. Briefly, 50 mg of P(DMC) was dissolved in 4 mL of DMF, and the solution was allowed to stir overnight. 40 mL of deionized water was then added dropwise and the mixture was maintained at 25 °C for 10 h. DMF was removed by dialysis (MWCO 3500) against deionized water to obtain the blank P(DMC) NPs and followed by lyophilization. DOX was encapsulated into P(DMC) NPs by a simple dialysis method. Before loading, DOX·HCl (5 mg, 0.0086 mmol) was stirred with TEA (2.4 µL, 0.0172 mmol) in DMSO (0.4 mL) for 2 h in the dark. 45 mg of P(DMC) was dissolved in 2 mL of DMF for 2 h, and then the DOX solution was added and the mixture was further stirred in the dark overnight. The solution was added lentamente into 40 mL of distilled water and stirred overnight. After that, the mixture was dialyzed (MWCO 3500) for 48 h to remove DMSO, DMF and unloaded DOX. After filtration, the obtained DOX-loaded P(DMC)NPs (P(DMC)@DOX NPs) were lyophilized in the dark. The control blank P(SA)NPs and DOX-loaded NPs (P(SA)@DOX NPs) were prepared in the same way. The amount of DOX was quantified via UV-vis spectrometer with the assistance of a calibration curve. DLC and DLE were obtained according to the following equations: DLC (wt%)= (weight of DOX in NPs/total weight of DOX-loaded NPs) × 100%; DLE (wt%)= (weight of DOX in NPs/total weight of DOX in feed) × 100%. 2.5. PH and reduction-triggered destabilization of P(DMC) NPs The size and size distribution changes of P(DMC) NPs in response to reduction and/or mildly


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acidic conditions were monitored by DLS measurements. The morphology change of P(DMC) NPs was followed by TEM observation. In general, lyophilized P(DMC) NPs were divided into six aliquots, which were adjusted into six different conditions: (i) PBS (pH 7.4), (ii) PBS (pH 6.5), (iii) acetate buffer (pH 5.0), (iv) PBS (pH 7.4, 10 µM GSH), (v) PBS (pH 7.4, 10 mM GSH) and (vi) acetate buffer (pH 5.0, 10 mM GSH). The final concentration of P(DMC) NPs was fixed at 0.5 mg mL-1. The solution was stirred at 37 °C. At predetermined time intervals, samples were taken for DLS and TEM measurements. 2.6. In vitro release of DOX/DMC Zeta potential measurements were performed to illuminate the DMC release indirectly. P(DMC) NPs were suspended in PBS 7.4, PBS 6.5, or acetate buffer pH 5.0 at 1 mg mL-1. P(SA) NPs were also dissolved in PBS as control. Subsequently, the copolymer solution was transferred to a dialysis bag (MWCO 3500) and dialyzed in 50 mL of the corresponding buffer at 37 °C with shaking. At predetermined time interval, samples were withdrawn and the zeta potentials were measured using a Zeta-Nanosizer. Afterwards, the samples were retransferred to the relevant dialysis bag. 5 mg of lyophilized P(DMC)@DOX NPs were dispersed in 5 mL of media in a dialysis tube (MWCO 3500), and dialyzed against 50 mL of corresponding media. The six different media conditions were (i) PBS (pH 7.4), (ii) PBS (pH 6.5), (iii) acetate buffer (pH 5.0), (iv) PBS (pH 7.4, 10 µM GSH), (v) PBS (pH 7.4, 10 mM GSH) and (vi) acetate buffer (pH 5.0, 10 mM GSH). At predetermined intervals, 1 mL of release media was withdrawn for analysis of the DOX concentration and replenished with the same amount of fresh media. The amount of released


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DOX was assayed by UV-vis spectrometer at 480 nm with the assistance of a standard curve. 2.7. In vitro studies Six cell lines, HepG2 (human hepatoma cells), HeLa (human cervical carcinoma cells), A549 (human non-small lung carcinoma cells), MCF-7 (human breast adenocarcinoma cells), SMMC-7721(human hepatoma cells) and DOX-resistant SMMC-7721cells were selected for cell tests. All cell lines were furnished by the Medical Department of Jilin University, China, and were incubated in Dulbecco’s modified Eagle’s medium (DMEM, GIBCO) at 37 °C in a 5% CO2 atmosphere. 2.7.1. Cellular uptake HeLa cells were cultured on the coverslips in 6-well plates (2 × 105cells per well) for 24 h. After adherence, the original medium was substituted with P(DMC)@DOX NPs at a final DOX concentration of 5.0 mg L-1 in PBS at pH 7.4 or 6.5 for a certain time. Then, cells were washed with cold PBS 7.4 and fixed with 4% formaldehyde for 30 min at room temperature, and the nuclei were stained with Hoechst 33258. The fluorescence images were obtained using confocal laser scanning microscopy (CLSM). To study the reduction-sensitivity of P(DMC)@DOX NPs, cells in six-well plate were cultured with 50 mM BSO for 12 h, or treated with 10 mM of GSH for 2 h. Cells without pretreatment were used as control. After that, cells were treated with free DOX or P(DMC)@DOX NPs for 4 h. Cells were washed with cold PBS 7.4 and fixed with 4% formaldehyde for 30 min at room temperature, and the nuclei were stained with Hoechst 33258. The fluorescence images were obtained using CLSM.


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For flow cytometry analysis, HeLa cells were seeded in 6-well plates (2 × 105 cells per well) with no coverslip. Cells were then treated with DOX or DOX-loaded NPs at different conditions. After trypsinization, the harvested cells were centrifuged at 1000 rpm for 5 min. The obtained cells were treated with three cycles of washing and centrifugation, and re-suspended in 0.5 mL PBS for flow cytometry analysis. 2.7.2. Cytotoxicity assay The in vitro cytotoxicity of different drug system (DOX, DMC, DOX + DMC, P(DMC) NPs, P(SA)@DOX NPs and P(DMC)@DOX) NPs) against HeLa cells under different reductive level was determined by a standard methyl tetrazolium (MTT) assay. In general, HeLa cells were seeded into 96-well plates at 8 × 103 or 4 × 103 cells per well in 100 µL of DMEM and further incubated for 12 h to adhere. Then, the predesigned cell groups were pretreated with 10 mM GSH for 2 h or 50 mM BSO for 12 h. After washing cells with cool DMEM, different drug groups were added with seven different DOX/DMC concentrations (for DOX: 0.156 ~ 10.0 µg mL-1; for DMC: 0.169 ~ 10.8 µg mL-1) and incubated for 24 or 48 h. HeLa cells incubated with blank DMEM were used as control. After predetermined incubation, MTT solution (20 µL, 5 mg mL-1) was added. Then, cells were cultured for another 4 h. The medium was aspirated, and the formazan generated by live cells was dissolved in 150 µL of DMSO. The relative cell viability was calculated by comparing the absorbance at 490 nm. The cytotoxicity of P(DMC)@DOX NPs against A549, MCF-7, HepG2, SMMC-7721 and DOX-resistant SMMC-7721 under different reductive level was also determined. The pH effect on cell viability of P(DMC)@DOX NPs was also evaluated against HeLa and


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A549 cells by MTT assay with the similar procedure as above. Briefly, after adherence for 24 h, cells were treated with 100 µL of fresh DMEM containing various concentrations of P(DMC)@DOX NPs at pH 7.4 or 6.8. After 4 h treatment, the solution was replaced with 200 µL of fresh DMEM medium at pH 7.4. Cells were then subjected to MTT assay after being cultured for another 24 or 48 h. 2.8. In vivo studies Female Kunming (KM) mice (6 ~ 8 weeks, weight ~ 25 g) were provided by Laboratory Animal Center, Jilin University (Changchun, China). All mice were maintained under required conditions and had free access to food and water throughout the experiments. A mouse hepatoma xenograft tumor model was generated by subcutaneous injection of H22 hepatoma cells (1 × 106) into the right flank of mice. All the in vivo study protocols were approved by the local institution review board and performed according to the Guidelines of the Committee on Animal Use and Care of Changchun Institute of Applied Chemistry, Chinese Academy of Sciences. 2.8.1. Bio-distribution When the tumor volume reached approximately 200 mm3, eighteen H22 tumor-bearing female KM mice were randomly separated into3 groups (6 mice in each group). Mice were administered intravenously with DOX·HCl (1.5 mg kg-1), P(SA)@DOX NPs (1.5 mg DOX kg-1) and P(DMC)@DOX NPs (1.5 mg DOX kg-1). At predetermined time intervals, three mice from each group were sacrificed. The major organs (heart, liver, spleen, lung and kidney) and tumor segment were excised. The DOX fluorescence of ex organs was imaged under the same parameters using a Maestro Imaging System (Cambridge Research & Instrumentation, Inc.,


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USA), in which the excitation time was fixed at 5 s. 2.8.2. Tumor inhibition and histological analysis Forty KM mice bearing H22 tumors were randomly divided into five groups (8 mice in each group) when the tumor volume was about 30 ~ 40 mm3, and this day was predetermined as day 1. Mice were treated with normal saline, DOX·HCl (3 mg kg-1), P(DMC) NPs (3.4 mg DMC kg-1), P(SA)@DOX NPs (3 mg DOX kg-1), and P(DMC)@DOX NPs (3.0 mg DOX kg-1, 3.4 mg DMC kg-1), respectively, via intravenous injection at day 1, 3, 6 and 9. Tumor length and width were measured with calipers, and the tumor volume was calculated using the following equation: tumor volume = length × width × width/2. The body weight and tumor volume of each mouse were measured every two or three days over a half month. At the end of treatment, mice were sacrificed and the tumors were excised for weighting and imaging. The main organs including heart, liver, spleen, lung and kidney were also collected, fixed in 4% paraformaldehyde overnight, then embedded in paraffin. The paraffin-embedded organs were cut at 5 mm thickness, and then stained with hematoxylin and eosin (H&E) to assess the histological alterations. 2.9. Statistical analysis The data of the experiments were expressed as means ± SD. Student’s test was used to analyze the statistical significance. Diffierences were considered statistically significant at a level of p < 0.05. 3. RESULTS AND DISCUSSION 3.1. Synthesis of multifunctional polymeric prodrug


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Since first reported by Sharpless et al. in 2002, CuAAC click reaction was extensively applied to synthesize multifarious functional polymers.40,41 Cystine, anatural amino acid with one disulfide bond in its structure, was selected to synthesize reduction-sensitive dialkyne monomer. To incorporate the charge-conversion property and anticancer activity into one system, anticancer drug DMC was reacted with the amino group of diazide monomer, and an acid-labile β-carboxylic amide group was formed simultaneously (Scheme 2). The dialkyned-cystine monomer was prepared following our previous work.38 The diazide monomer was elaborately characterized by 1H NMR, 13C NMR, FT-IR, MALDI-TOF (Figure S1). All peaks in the 1H and 13

C NMR spectra were well assigned, and the peak at 2101 cm−1 in the FT-IR spectrum of

DMC-conjugated diazide monomer was attributed to N=N=N. The chemical characterization confirmed the successful synthesis of DMC-conjugated diazide monomer with high purity.


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Scheme 2 Synthetic routes of the clickable monomers, and ABA-type triblock copolymers P(DMC) and P(SA). The ABA-type triblock multifunctional polymeric prodrug P(DMC), mPEG5k-b-poly (disulfide-alt-demethylcantharidin)-b-mPEG5k could be synthesized by step polymerization of reduction-sensitive dialkyne monomer and pH-sensitive DMC-conjugated diazide monomer with CuCl/PMDETA as catalysis and mPEG5k-N3as blocking segments in a controlled way. Typical 1

H NMR spectra of P(DMC) is shown in Figure S2. The resonances at 3.51, 1.37, 8.33 and 1.61


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ppm were ascribed to the methylene or methyl protons of PEG, BOC, triazole and DMC segments, respectively. The degree of polymerization (DP) of P(DMC) was calculated to be 8.7, by comparing the integration of methylene peak of PEG (−CH2−CH2−) with that of methylene peak of DMC unit (−CH2−CH2−). GPC analysis exhibited that P(DMC) had a single peak (Mn = 4.08 × 104 g mol-1) and narrow molecular weight distribution (PDI = 1.21) (Figure S3). Furthermore, disappearance of azide peak at 2101 cm-1 in the FT-IR spectrum also illustrated the complete elimination of superfluous mPEG5k-N3 (Figure S4). All the data demonstrated that the monomers and polymeric prodrug were successfully synthesized. The DLC of DMC was calculated to be 8.6%, which could be also easy regulated by adjusting DP and hydrophilic/hydrophobic block ratio of the polymeric prodrug. As a control copolymer, P(SA) without charge-conversion property and anticancer activity was also prepared in a similar way, in which succinic anhydride instead of DMC was used to synthesize the diazide monomer (Figure S4-S6). 3.2. Preparation and characterization of polymeric prodrug nanoparticles Amphiphilic ABA-type triblock copolymers, P(DMC) and P(SA), with two hydrophilic blocks and one hydrophobic block, could self-assemble into micelle-type nanoparticles in aqueous solution. Critical aggregation concentration (CAC) is one of the significant parameters to reflect the self-assembly stability of micelle-type nanoparticles. As shown in Figure S7, the CAC values of P(DMC) and P(SA) were calculated to be 4.816 mg L-1and 4.912 mg L-1, respectively. The low CAC values demonstrated great stability of the aggregates against dilution, propitious to intravenous administration. DLS was also used to investigate the size and size distribution. For


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blank P(DMC) NPs, the average size was about 102 nm and the size distribution was considerable narrow. After DOX encapsulation, the size and size distribution of P(DMC)@DOX NPs (DOX DLC: 7.9%; DLE: 79.0%) slightly increased (Figure 1). Both P(DMC) and P(DMC)@DOX NPs showed clear spherical morphology by TEM observation, and the average diameters were smaller than DLS results, which may be due to the shrinkage of PEG shell. Similar DLS and TEM results of P(SA) NPs and P(SA)@DOX NPs (DOX DLC: 7.1%; DLE: 74.6%) were also shown.

Figure 1. Typical TEM images and hydrodynamic radius distribution (Rh) of (a) P(DMC) NPs, (b) P(DMC)@DOX NPs, (c) P(SA) NPs and (d) P(SA)@DOX NPs (scale bar: 500 nm). 3.3. Acidic PH and reduction dual-sensitivity of P(DMC) NPs The significant characteristics of P(DMC) were that reduction-sensitive disulfide groups were arranged repeatedly in the backbone of hydrophobic segment, and β-carboxylic amide groups were regularly localized on the side chains. It is well known that β-carboxylic amide structure is quite stable at basic and neutral pH, but unstable at acidic pH which hydrolyzes into corresponding dicarboxylic acid/anhydride and amine.42,43 The unique characters of disulfide and β-carboxylic amide were therefore applied for our designed polymeric prodrug NPs to own pH and reduction dual-sensitivity.


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Figure 2. Average diameter and size distribution (PDI) of P(DMC) NPs as a function of time under different (a) pH and (b) reductive level determined by DLS (solid line: average diameter; dot line: PDI). Kinetic stability of P(DMC) NPs under different pH was investigated by DLS and TEM measurements. Average diameter and polydispersity index (PDI) were plotted as a function of time. As shown in Figure 2a, at pH 7.4, the size and PDI of P(DMC) NPs were about 99 nm and 0.1, respectively. The values did not change during the entire testing time, indicating the stability of P(DMC) NPs at pH 7.4, beneficial to long circulation. When pH was adjusted to 6.5, the average diameter of P(DMC) NPs immediately increased to 156 nm, which could be attributed to the loss of colloidal stability and aggregation after surface charge was partially reversed. Though DMC were partially released from the side chain of polymer, the structure of the polymer backbone was not affected and no obvious changes were observed for the average diameter and the PDI with time. If pH was further regulated to 5.0, the size and distribution both increased sharply, showing clear pH-induced nanoparticle dissociation. Distinct concentrations of reductive GSH (10 µM and 10 mM) were further utilized to mimic the physiological and intracellular


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reductive levels for reduction-sensitivity evaluation. As can be seen from Figure 2b, only slight change was found for P(DMC) NPs in the presence of 10 µM of GSH at pH 7.4, indicating the high stability under circulation. However, the size and distribution had tremendous variety in the presence of 10 mM GSH at pH 7.4/5.0 with time. The degradation and re-aggregation of NPs developed immediately, with bimodal and broad size distribution patterns showed in DLS (Figure 2 and Figure S8). TEM images of P(DMC) NPs under different conditions at predetermined time point (24 h) were also shown in Figure 3. At pH 7.4 without GSH, P(DMC) NPs showed stable and uniform spherical shape. When pH was adjusted to acidic (6.5 or 5.0), irregular morphology of P(DMC) NPs was found. Especially, in the presence of 10 mM GSH and at pH 5.0, the size and morphology of P(DMC) NPs had huge change, and some smaller NPs formed

by

hydrophobic

fragments

with

high

contrast

appeared,

suggesting

the

pH/reduction-induced disintegration. All the results were consistent with the hypothesis of the pH/reduction dual-sensitive characteristics of P(DMC) NPs.


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Figure 3. Typical TEM images of P(DMC) NPs suspended in different solutions for 24 h: (a) pH 7.4; (b) pH 6.5; (c) pH 5.0; (d) pH 7.4 + 10 µM GSH; (e) pH 7.4 + 10 mM GSH; (f) pH 5.0 + 10 mM GSH (white scale bar: 500 nm, blue scale bar: 1 µm). 3.4. Charge-conversion and triggered drug release of P(DMC) NPs In our case, an anticancer drug, DMC was introduced onto the side chains of copolymer to produce β-carboxylic amide structure.44,45 Hence, P(DMC) is actually the polymeric prodrug of DMC. Only when the β-carboxylic amide groups are hydrolyzed under slightly tumor extracellular acidic environment can the amine groups on side chains of polymeric prodrug be recovered, inducing the charge-conversion of P(DMC) NPs for enhanced cellular internalization and DMC release for anticancer ability. The determination of DMC content was comparatively arduous. Consequently, zeta potential


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measurement was utilized to indicate the charge-conversion and DMC release. As shown in Figure 4a, P(DMC) NPs had an initial zeta potential of about − 20 mV at pH 7.4, which changed little over 48 h, indicating the difficult hydrolysis of DMC from P(DMC) NPs at pH 7.4. However, P(DMC) NPs underwent a clear charge reversal from − 20 to + 5.54 mV at pH 6.5 in 48 h. If pH was further declined to 5.0, zeta potential of P(DMC) NPs sharply increased to positive charge within 9 h, showing the efficient hydrolysis of DMC. However, control P(SA) NPs did not hydrolyze clearly in the acidic condition, showing strong negative charge during the test time with slight deviation. Taking the negatively charged cell membranes into consideration, the negative-to-positive charge-conversion ability of P(DMC)NPs at the slightly acidic tumor extracellular environment would be beneficial to their internalization by tumor cells. After DOX encapsulation, the DOX release behavior of P(DMC)@DOX NPs was determined using a dialysis method at 37 °C in different buffer solution (Figure 4b). As expected, P(DMC)@DOX NPs exhibited relative stability at pH 7.4 with no more than 12% of DOX release after 72 h. In addition, there was no obvious change of DOX release in the presence of 10 µM GSH, which could be propitious to reduce the premature release during blood circulation. The 72 h cumulative release of DOX slightly increased from 11.4% at pH 7.4 to 19.2% at pH 6.5. As the pH declined to 5.0, more than 49.9% of DOX were released within the same period, indicating that DOX release from P(DMC)@DOX NPs was sensitive to the endo/lysosomal pH. The accelerated DOX release might be explained by the increased solubility of DOX and also the charge-conversion induced instability of NPs in acid conditions. When P(DMC)@DOX NPs were incubated in the presence of 10 mM GSH at pH 7.4 or 5.0, over 53% and 85% of DOX


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were released, respectively. The results demonstrated that P(DMC)@DOX NPs had excellent pH and reduction dual-sensitivity towards intracellular acid and GSH stimuli simultaneously for efficient drug delivery into cancer cells.

Figure 4. (a) Zeta potentials of P(DMC) NPs and P(SA) NPs, and (b) DOX release profiles of P(DMC)@DOX NPs under different conditions. 3.5. Cellular internalization Previous studies have demonstrated that polymeric NPs can be internalized by cancer cells via energy-using endocytosis, much different with free drugs through passive diffusion.46 Therefore, cellular internalization of P(DMC)@DOX NPs against HeLa cells at different temperature (4 °C and 37 °C) was monitored by CLSM (Figure S9). The uptake was temperature-depended, indicating the endocytosis-mediated internalization of P(DMC)@DOX NPs. To illuminate whether the charge-conversion ability of P(DMC)@DOX NPs at tumor extracellular pH would affect endocytosis, the cellular internalization after 2 h incubation both at pH 6.5 and 7.4 were determined (Figure 5). Compared to pH 7.4, extremely stronger DOX fluorescence was observed in cells at pH 6.5. DOX was mainly aggregated around nuclei at pH 7.4, while strong


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fluorescence was observed in the verge of nuclei at pH 6.5, and the depth could be clearly observed. To further elucidate the effect of charge-conversion ability on cellular uptake, the cellular accumulation of P(DMC)@DOX NPs and P(SA)@DOX NPs at pH 7.4/6.5 was quantitatively conducted by flow cytometry. As shown in Figure S10, compared with cells incubated with P(SA)@DOX NPs, cells treated with P(DMC)@DOX NPs at pH 6.5 for 2 h displayed a distinct right shift of fluorescence than those at pH 7.4. It is reasonable that P(DMC)@DOX NPs would perform surface negative-to-positive charge conversion at pH 6.5, resulting in promoted charge interaction with cell membrane. The results demonstrated that P(DMC)@DOX NPs with charge-conversion ability at tumor extracellular pH exhibited greatly enhanced cellular internalization.

Figure 5. CLSM images of HeLa cells cultured with P(DMC)@DOX NPs at different pH (6.5 or 7.4) for 2 h (scale bar: 50 µm). The cellular internalization behavior of P(DMC)@DOX NPs in the presence or absence of GSH/BSO at pH 7.4 for 4 h was further investigated (Figure 6). HeLa cells were pretreated with 50 mM BSO to inhibit the intracellular synthesis of GSH, or with 10 mM of GSH to enhance the intracellular reductive concentration, respectively. As expected, different from the rapid diffusion


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of free DOX into nuclei, DOX fluorescence was mainly observed around rather than in nuclei in the case of P(DMC)@DOX NPs. For BSO-pretreated cells, the fluorescent intensity in cells was weakened and the DOX localization showed no significant change. In stark contrast to BSO-pretreated cells, it’s notable that the intensity was greatly enhanced for GSH-pretreated cells and strong fluorescence was detected in nuclei. This is in agreement with the flow cytometry results shown in Figure S11. All the results revealed that the enhanced DOX fluorescent intensity in the GSH-pretreated cells should ascribe to the improved intracellular release of DOX beneficial from the reduction-induced NPs degradation.

Figure 6. CLSM images of HeLa cells incubated with DOX or P(DMC)@DOX NPs under different condition at pH7.4 for 4 h (scale bar: 50 µm). 3.6. Cytotoxicity evaluation The above studies illustrated that as pH decreased, zeta potential of P(DMC) NPs would perform negative-to-positive conversion due to the hydrolysis of DMC-conjugated β-carboxylic amide, resulting in swelling and decomposition of NPs, enhanced cellular uptake and accelerated intracellular drug release.36 The anticancer activity of DMC has been extensively investigated against a diversity of malignancies. In this work, DMC was not only used to introduce


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β-carboxylic amide, but also acted as an anticancer drug for co-delivery with the encapsulated DOX. Hence, in vitro cytotoxicity of P(DMC)@DOX NPs under different pH were investigated by MTT assay. HeLa cells were cultured with P(DMC)@DOX NPs at pH 7.4 or 6.5 for 4 h, then the medium was replaced with complete DMEM for further incubation. Cells cultured without drug were used as positive control. As depicted in Figure 7a, P(DMC)@DOX NPs exhibited the obviously improved cytotoxicity at DOX concentration higher than 2.5 µg mL-1 at pH 6.5 compared to that at pH 7.4, due to the enhanced cellular internalization and DMC release through mild acidic pH-triggered charge-conversion of P(DMC)@DOX NPs. Similar result was also found in A549 cells (Figure S12). Cytotoxicity of DOX, DMC, P(DMC)NPs, P(SA)@DOX NPs and P(DMC)@DOX NPs under different reductive level against HeLa cells were also investigated. BSO and 10 mM GSH pretreatment were applied as well. As shown in Figure 7b, free DOX, DOX + DMC, and GSH-pretreated P(DMC)@DOX NPs groups had equivalent and highest anticancer activity. For other groups, the IC50 values increased in the following order: P(DMC)@DOX NPs < BSO-pretreated P(DMC)@DOX NPs < P(SA)@DOX NPs < DMC < P(DMC) NPs (Table S1). Although showed the lowest anticancer activity, P(DMC) NPs (without any pretreatment) could inhibit the cell proliferation to some extent, probably due to the released DMC derivative and the charge-conversion property compared with P(SA) NPs (Figure S13). For the same reason, P(DMC)@DOX NPs exhibited higher anticancer activity than that of P(SA)@DOX NPs. Similar results were also observed in A549, MCF-7, HepG2 and SMMC-7721 cell lines (Figure 7c, S12 and S14). All the data proved that our multifunctional polymeric prodrug NPs were a promising


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controlled co-delivery system for combination chemotherapy. It’s well known that multidrug resistance (MDR) remains the major impediment for successful chemotherapy.47 It was reported that DMC could reverse cisplatin resistance by PP2A inhibition, 48,49

and overcome MDR to DOX by regulating sonic hedgehog signaling.50 Here, the anticancer

activity of P(DMC)@DOX NPs against DOX-resistant SMMC-7721 cells was further investigated. As shown in Figure 7c-7d, DOX-resistant SMMC-7721 cells showed notably high resistance to DOX (IC50 = 67.2 µg/mL), and the resistant factor (RF) was calculated to be 47 after 24 hour’s incubation (Table S1, Figure S15). However, a simple combination of free DOX and DMC could significantly decrease the IC50 value of DOX to 23.5 µg mL-1, indicating the strong effect of DMC to improve DOX sensitivity against DOX-resistant cells. While in the case of P(DMC)@DOX NPs and GSH-pretreated P(DMC)@DOX NPs groups, the IC50 values were further decreased to 15.4 and 10.6 µg mL-1, respectively, which were comparable to the IC50 value for free DOX against SMMC-7721 cells (1.4 µg mL-1). With the high cell proliferation inhibition activity against DOX-resistant SMMC-7721 cells, the new multifunctional P(DMC)@DOX NPs provide a potential platform for more efficient treatment of tumors.


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Figure 7. (a) Cell viability of P(DMC)@DOX NPs against HeLa cells at different pH for 24 h; Cell viability curves of various drug systems against (b) HeLa cells, (c) SMMC-7721 cells and (d) DOX-resistant SMMC-7721 cells for 24 h. 3.7. Ex vivo DOX fluorescence imaging Polymeric drug delivery vehicles have the ability to improve drug pharmacokinetics, bio-distribution and tissue-specific accumulation. To further demonstrate the enhanced tumor accumulation of P(DMC)@DOX NPs by virtue of charge-conversion ability, the ex vivo bio-distribution was investigated by DOX fluorescence imaging. At predetermined time interval


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(6 h, 24 h), mice injected with 1.5 mg kg-1 of DOX, P(SA)@DOX NPs or P(DMC)@DOXNPs were sacrificed. The isolated organs involved heart, liver, spleen, lung, kidney and tumor components. The photographs of the excised organs were list in Figure 8. It was found that liver showed stronger DOX fluorescence for all groups, especially for NPs groups, which might due to more blood volumes/gm of liver as well as active phagocytosis by macrophages in the liver.51,52 For mice treated with free DOX, tumor exhibited very strong DOX fluorescence at 6 h post injection, but the intensity weakened obviously with time prolonged to 24 h. In contrast, for DOX-loaded NPs, the fluorescence intensity in tumor remained stable with time, attributed to the EPR effect of nanoparticles. Interestingly, we also found that P(DMC)@DOX NPs showed much higher tumor accumulation than P(SA)@DOX NPs, benefit from the negative-to-positive charge-conversion ability of our polymeric prodrug NPs for enhanced cellular internalization and accelerated/sufficient DOX release after tumor accumulation.

Figure 8. Ex vivo DOX fluorescence imaging of H22 tumor-bearing KM mice after 6 h and 24 h


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post-injection of DOX, P(SA)@DOX NPs and P(DMC)@DOX NPs (a, heart; b, liver; c, spleen; d, lung; e, kidney; f, tumor). 3.8. In vivo antitumor efficacy and safety Finally, based on the above results, the in vivo antitumor efficacy of P(DMC)@DOXNPs was evaluated on H22 hepatoma xenograft tumor-bearing female KM mice. Mice were treated with saline, free DOX·HCl (3.0 mg kg-1), P(DMC) NPs (3.4 mg DMC kg-1), P(SA)@DOX NPs (3.0 mg DOX kg-1) and P(DMC)@DOX NPs (3.0 mg DOX kg-1, 3.4 mg DMC kg-1), accordingly. The tumor volumes and bodyweights were measured every two or three days. The treatment with P(SA)@DOX NPs and P(DMC)@DOX NPs did not result in any obvious body weight loss (Figure 9b), demonstrating the well-tolerated and reduced systemic toxicity of drug-loaded NPs for potential clinical utility. As shown in Figure 9a, the tumor volume of saline treated group increased extra ordinarily fast, and the average tumor volume reached to approximately 3000 mm3 within 17 days. In comparison to the negative control group, there was a slight inhibition efficacy for mice treated with P(DMC) NPs, and the average tumor volume was about 1700 mm3 at the end of treatment. It was found that mice treated with free DOX had good suppression on tumor growth only in the first 10 days, after that, tumor growth was out of control. Instead, DOX-loaded NPs groups showed long-term inhibition efficacy on tumor growth, and P(DMC)@DOXNPs demonstrated the best antitumor effect and the tumor growth was completely restricted during the test period. The holistic tumor photograph after excision was visible in Figure 9c. It is notable that at the end of treatment, the tumor volume of saline, P(DMC)


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NPs, DOX and P(SA)@DOX NPs groups increased by 94.2-, 57.8-, 21.8- and 6.5-fold, respectively, whereas P(DMC)@DOX NPs group increased by only 2.4-fold. The significantly improved antitumor efficiency of P(DMC)@DOX NPs should be ascribed to the efficient co-delivery of DMC and DOX into tumor site via EPR effect, charge-conversion induced tumor cellular uptake enhancement, and sufficient intracellular drug release.

Figure 9. (a) Tumor growth curves and (b) relative body weight changes of H22 tumor-bearing KM mice after different treatment. (c) Pictures of excised tumors on day 17 (dividing rule: cm). Data are expressed as mean ± standard deviation, n = 8, *P < 0.05. Long-term toxicity is a major concern for in vivo application of chemotherapy. The histological analysis of fixed tissues was showed in Figure 10. No obvious abnormality was


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found in spleen and lung for all groups. However, compared with saline group, DOX formulation-treated groups showed pathological changes more or less in heart, liver and kidney. Particularly, free DOX displayed noticeable signals of damage in the heart, with the irregularly arranged myocardial cells and necrosis of muscle fibers in the cardiac tissues. While the treatment with P(SA)@DOX NPs and P(DMC)@DOX NPs obviously reduced the blight of heart, demonstrating the reduced systemic toxicity for potential clinical application.

Figure 10. Histological analysis of tissues with H&E after treatments with saline, P(DMC) NPs, free DOX, P(SA)@DOX NPs and P(DMC)@DOX NPs. 4. CONCLUSION


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In the present study, we demonstrated a pH and reduction dual-sensitive charge-conversional polymeric prodrug strategy. Disulfide group and DMC-conjugated β-carboxylic amide group were repeatedly and regularly introduced into the polymer chain simultaneously via facile CuAAC click polymerization. The obtained polymeric prodrug, P(DMC), was further used to encapsulate DOX for efficient co-delivery with DMC. The surface charge of P(DMC)@DOX NPs could perform negative to positive conversion in the slightly acidic tumor microenvironment. Under acidic pH and 10 mM GSH conditions, P(DMC)@DOX NPs showed remarkably enhanced cellular internalization and cytotoxicity in vitro, especially against DOX-resistant SMMC-7721 cells. The in vivo studies on H22 tumor bearing KM mice illustrated that P(DMC)@DOX NPs exhibited higher tumor accumulation and improved antitumor efficiency compared to P(SA)@DOX NPs with one drug and without charge-conversion ability. In summary, P(DMC) NPs could be smartly “switched on” in response to certain stimuli or phenotypes in tumor tissue/cell microenvironments, thereby for efficient co-delivery and controlled release of DOX and DMC. The desired multifunctional polymeric prodrug strategy brings a new opportunity for cancer chemotherapy. ASSOCIATED CONTENT Supporting Information. Additional experimental protocols, characterization data and in vitro results are available at free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author


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* Dongfang Zhou: Tel: +86-431-85262538; E-mail: [email protected] Yubin Huang: Tel: +86-431-85262769; E-mail: [email protected]. ACKNOWLEDGEMENTS The authors would like to thank the financial support from the National Natural Science Foundation of China (No.51403198 and 51573069), and Jilin Provincial Science and Technology Department (No.20150520019JH).

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(8) Zhang, Z.; Chen, X.; Chen, L.; Yu, S.; Cao, Y.; He, C.; Chen, X., ACS Applied Materials & Interfaces 2013, 5, 10760-10766. (9) Zhao, Z.; Yao, X.; Zhang, Z.; Chen, L.; He, C.; Chen, X., Macromolecular Bioscience 2014, 14, 1609-1618. (10) Ma, N.; Li, Y.; Xu, H. P.; Wang, Z. Q.; Zhang, X., Journal of the American Chemical Society 2010, 132, 442-443. (11) Li, J.; Qu, X.; Payne, G. F.; Zhang, C.; Zhang, Y.; Li, J.; Ren, J.; Hong, H.; Liu, C., Advanced Functional Materials 2015, 25, 1404-1417. (12) Xu, W.; Ding, J.; Xiao, C.; Li, L.; Zhuang, X.; Chen, X., Biomaterials 2015, 54, 72-86. (13) de la Rica, R.; Aili, D.; Stevens, M. M., Advanced Drug Delivery Reviews 2012, 64, 967-978. (14) de Gracia Lux, C.; Joshi-Barr, S.; Nguyen, T.; Mahmoud, E.; Schopf, E.; Fomina, N.; Almutairi, A., Journal of the American Chemical Society 2012, 134, 15758-15764. (15) Liu, J.; Pang, Y.; Chen, J.; Huang, P.; Huang, W.; Zhu, X.; Yan, D., Biomaterials 2012, 33, 7765-7774. (16) Cheng, R.; Meng, F.; Deng, C.; Klok, H.-A.; Zhong, Z., Biomaterials 2013, 34, 3647-3657. (17) Jhaveri, A.; Deshpande, P.; Torchilin, V., Journal of Controlled Release 2014, 190, 352-370. (18) Li, Y.; Xiao, K.; Zhu, W.; Deng, W.; Lam, K. S., Advanced Drug Delivery Reviews 2014, 66, 58-73. (19) Chen, P.; Qiu, M.; Deng, C.; Meng, F.; Zhang, J.; Cheng, R.; Zhong, Z., Biomacromolecules 2015, 16, 1322-1330.


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