Surface Functionalization of Polymeric Nanoparticles with Umbilical

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Biological and Medical Applications of Materials and Interfaces

Surface Functionalization of Polymeric Nanoparticles with Umbilical CordDerived Mesenchymal Stem Cell Membrane for Tumor-Targeted Therapy Na Yang, Yanping Ding, Yinlong Zhang, Bin Wang, Xiao Zhao, Keman Cheng, Yixin Huang, Mohammad Taleb, Jing Zhao, Wen-Fei Dong, Lirong Zhang, and Guangjun Nie ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b05363 • Publication Date (Web): 15 Jun 2018 Downloaded from http://pubs.acs.org on June 15, 2018

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Surface Functionalization of Polymeric Nanoparticles with Umbilical Cord-Derived Mesenchymal Stem Cell Membrane for Tumor-Targeted Therapy ⊥

⊥

Na Yang†‡§#, Yanping Ding‡ *, Yinlong Zhang‡$, Bin Wang‡ , Xiao Zhao‡$, Keman Cheng‡, Yixin Huang‡⊥, Mohammad Taleb‡⊥, Jing Zhao†, Wen-Fei Dong§*, Lirong Zhang$, Guangjun Nie‡⊥* †

School of Life Sciences, Shanghai University, No. 333 Nanchen Road, Baoshan District,

Shanghai 200444, China ‡

CAS Key Laboratory for Biomedical Effects of Nanomaterials & Nanosafety, CAS Center for

Excellence in Nanoscience, National Center for Nanoscience and Technology (NCNST), 11 Beiyitiao, Zhongguancun, Beijing 100190, China §

CAS Key Laboratory of Bio-Medical Diagnostics, Suzhou Institute of Biomedical Engineering

and Technology (SIBET), No.88 Keling Road, Suzhou New District 215163, Jiangsu Province, China ⊥

$

University of Chinese Academy of Sciences, Beijing 100049, China

School of Basic Medical Sciences, Zhengzhou University, Zhengzhou, Henan, 450001, China

KEYWORDS: umbilical cord mesenchymal stem cell, plasma membrane, biomimetic nanoparticle, targeted drug delivery, cancer therapy


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ABSTRACT Multiple cell plasma membranes have been utilized for surface functionalization of synthetic nanomaterials and construction of biomimetic drug delivery systems for cancer treatment. The natural characters and facile isolation of original cells facilitate the biomedical applications of plasma membranes in functionalizing nanocarriers. Human umbilical cord-derived mesenchymal stem cells (MSC) have been identified to show tropism towards malignant lesions and have great advantages in ease of acquisition, low immunogenicity, and high proliferative ability. Here we developed a poly(lactic-co-glycolic acid) (PLGA) nanoparticle with a layer of plasma membrane from umbilical cord MSC coating on the surface for tumor-targeted delivery of chemotherapy. Functionalization of MSC plasma membrane significantly enhanced the cellular uptake efficiency of PLGA nanoparticles, the tumor cell killing efficacy of PLGA-encapsulated doxorubicin, and most importantly the tumor-targeting and accumulation of the nanoparticles. As a result, this MSC-mimicking nanoformulation led to remarkable tumor growth inhibition and induced obvious apoptosis within tumor lesions. This study for the first time demonstrated the great potential of umbilical cord MSC plasma membranes in functionalizing nanocarriers with inherent tumor-homing features, and the high feasibility of such biomimetic nanoformulations in cancer therapy.


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INTRODUCTION Functional nanomaterials have been used for construction of various drug delivery systems for cancer therapy.1-2 The advantages of cancer nanomedicines lie in tumor-targeting and accumulation, tumor-responsive release of drugs, increase in drug stability, and reduction of adverse effects.3-4 Numerous studies have focused on improving the tumor-targeting efficiency and avoiding non-specific elimination of nanocarriers,5 mainly by surface modification of functional moieties such as peptide ligands or polyethylene glycol (PEG).6-8 These strategies usually require complicated chemical synthesis and are difficult for large-scale production. Cellular plasma membranes participate in many vital biological processes through proteins embedded in the phospholipid bilayer. As natural biomaterials, plasma membranes show great biocompatibility and innately possess various features of the specific cell origin. Multiple cell plasma membranes have been utilized to generate biomimetic nanocarriers for treatment of specific diseases.9 An erythrocyte membrane-coated polymer nanoparticle has been reported to inherit the long-circulating property of red blood cells.10 Silicon nanoparticles modified with leukocyte cell membrane have been found to target inflammation region because the active adhesion molecule lymphocyte function-associated antigen 1 (LFA-1) can bind intercellular cell adhesion molecule-1 (ICAM-1) on inflamed endothelium.11 Based on the interaction between platelet and tumor cells, platelet membrane has been used for engineering tumor-targeting nanoparticles for multiple myeloma and head and neck squamous cell carcinoma.12, 13 Cancer cell-derived membranes have been shown to functionalize nanocarriers with tumor-homing ability,14 and can provide tumor antigen to serve as a cancer vaccine.15, 16


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Mesenchymal stem cells (MSCs) have been reported to show tropism towards inflammatory or malignant tissues in many experimental animal models, such as glioma, colon cancer, breast cancer and hepatocellular carcinoma (HCC).17-20 The underlying mechanism is complicated, mainly involving chemokine-receptor interaction and adhesion to endothelium.21 Tumor cells and tumor-associated stromal cells have been identified to produce a large amount of chemokines, including vascular endothelial growth factor (VEGF), platelet derived growth factor (PDGF), stromal cell-derived factor-1 (SDF-1), and transforming growth factor-β (TGF-β), that can attract MSCs with corresponding receptors expressed on the cell surface.22 The tumorhoming feature of MSCs also depends on their interaction with endothelial cells through adhesion molecules vascular cell adhesion molecule (VCAM) and intercellular cell adhesion molecule-1 (ICAM).23, 24 Due to the natural tumor-targeting capacity and low immunogenicity, MSCs-derived from bone marrow and adipose tissue have been used for delivery of therapeutics to tumors.25,

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However, MSCs play intricate roles in tumor progression and metastasis.27

Utilization of MSCs for cancer treatment may provoke a risk, such as inducing metastasis. Compared with MSCs, MSC plasma membrane is regarded as a safer drug delivery platform, and bone marrow MSC membrane has been used for drug delivery in prostate cancer and cervical cancer.28, 29 Nevertheless, the possible injury or severe side effects during bone marrow isolation is an obstacle to the potential broad application of bone marrow MSC membrane. Recently, human umbilical cord-derived MSCs gain great attention in clinical application due to the ease of acquisition, expansion, and maintaining stemness, providing a desirable source of MSC plasma membranes.30,

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Meanwhile, clinical application of MSCs for treatment of graft-versus-host

disease has been reported to be irrespective of tissue type matching,32 further suggesting the high potential of MSCs and MSC plasma membranes used in clinical translation. However, the


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performance of umbilical cord MSC-coated nanoparticles in cancer treatment has not been reported yet. Inspired by the critical roles of umbilical cord-derived MSCs and plasma membrane-coated nanoparticles, we developed umbilical cord-derived MSC plasma membrane-coated polymeric nanoparticles for tumor-targeted delivery of chemotherapy. The functional nanocarrier exhibited high tumor accumulation efficiency and potentiated the antitumor effects of chemotherapeutic drugs, with minimal adverse effects. This study for the first time demonstrated the potential of plasma membranes from umbilical cord MSCs in functionalizing synthetic nanomaterials, and provided an alternative strategy for engineering tumor-targeted drug delivery system with natural and multiple targeting moieties.

MATERIALS AND METHODS Cells and materials. Human mesenchymal stem cells (MSCs) were kindly provided by Beijing Cellonis Biotechnology Corporation (Beijing, China) and were isolated from human umbilical cord according to a previous report.33 Human hepatocellular carcinoma cell line MHCC97H and human umbilical vein endothelial cell line (HUVEC) were purchased from Jennio Biotech (Guangzhou, China). Phosphate buffered saline (PBS), RPMI 1640 medium and fetal bovine serum were purchased from Wisent (St. Bruno, Canada). Mesenchymal stem cell growth medium was a BulletKit medium (cat. 00190632) including mesenchymal basal medium and the necessary supplements for MSC proliferation, and was purchased from Lonza (USA). Phosphatase and protease inhibitor cocktail was purchased from Roche (Shanghai, China). BCA protein assay kit was from Thermo Fisher Scientific (Waltham, USA). Lecithin was from Lipoid


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GmbH (Germany). Cholesterol was purchased from J&K (China). DSPE-PEG2000 was from Nanocs (USA). Rhodamine B, PKH 67 and Indocyanine green (ICG) were purchased from Sigma (USA). Lyso-sensor Green DND 189 was purchased from Invitrogen (USA). Cell counting kit-8 (CCK-8) was obtained from Dojindo Molecular Technologies (Kumamoto, Japan). Poly(lactide-co-glycolide) (PLGA, molar ratio of D, L-lactic to glycolic acid, 75: 25; molecular weight, 15 kD) was purchased from Jinan Daigang Biomaterial (Shandong, China). Doxorubicin hydrochloride was purchased from Beijing Mesochem Technology (Beijing, China). Anti-caspase 3 (cat. 9662S) and anti-cleaved caspase 3 (cat. 9664S) were from Cell Signaling Technology (USA). Anti-CXCR4 (cat. 60042) was from ProteinTech (USA). Anti-Na+/K+ ATPase (cat. ab76020) was from Abcam (USA). Anti-GAPDH and anti-β-actin were from Beyotime (China). Cell culture. MHCC97H cells and HUVECs were cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum. MSCs were maintained in Lonza TheraPEAK™ mesenchymal stem cell growth medium and cells between 3-7 passages were used. All cells were maintained at 37 oC in a humidified incubator with 5% CO2. Isolation of MSC plasma membrane. The MSC plasma membrane was isolated according to a previous study.34 When MSCs were 90% confluent, cells were washed twice using ice-cold PBS buffer and were harvested in 3 mL of buffer (pH 7.8) containing 250 mM sucrose, 20 mM tricine, and 1 mM EDTA. After cell suspension was centrifuged at 1000 g for 5 min, cell pellet was collected, homogenized in 1 mL of buffer (pH 7.4) containing 225 mM mannitol, 75 mM sucrose, 0.5% BSA, 0.5 mM EGTA, 30 mM Tris, and phosphatase and protease inhibitor cocktail, and further centrifuged at 10,000 g for 10 min at 4 oC. The supernatant was then ultracentrifuged at 100,000g for 60 min at 4 oC. The plasma membrane pellet was resuspended


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in buffer (pH 6.0) containing 5 mM Bis-Tris and 0.2 mM EDTA and stored at -80 oC for further experiment. The protein concentration of purified plasma membrane was quantified using a BCA protein assay kit. Preparation of Dox-loaded PLGA nanoparticles (NP-Dox). PLGA nanoparticles were prepared using double-emulsion method according to a previous study.35 In brief, PLGA was dissolved in dichloromethane at a concentration of 10 mg/mL. Then 1 mL of PLGA solution was mixed with 0.2 mL of Dox aqueous solution (2 mg/mL). The mixture was emulsified under ultrasonic (100 W) for 2 min, followed by mixing with 2 mL of 2% sodium cholate solution and ultrasonication for 2 min. The emulsion was dropwise added with 10 mL of 0.5% sodium cholate solution and stirred for 1 h at room temperature. After rotary evaporation, NP-Dox harvested by centrifugation of the emulsion at 10,000g for 15 min and washed twice using distilled water. Construction of PLGA nanoparticles coated with plasma membrane or liposome. Coating of plasma membrane onto PLGA nanoparticles was performed according to previous reports.36 In brief, 500 µL of PLGA nanoparticles (2 mg/mL) was mixed with 40 µL of MSC plasma membrane solution (protein concentration, 5 mg/mL), and was ultrasonicated (100 W) for 5 min. The mixture was centrifuged at 10,000 g for 10 min and the pellet was plasma membrane-coated PLGA nanoparticles (PM-NPs). To prepare liposome-coated PLGA nanoparticles (Lipo-NPs), liposome was first prepared by the thin-film hydration method. Briefly, 3 mg of lectin, 1.5 mg of cholesterol and 0.5 mg of DSPE-PEG2000 were dissolved in 1 mL of chloroform in a round bottom flask. After rotary evaporation, a thin phospholipid film was formed at the flask bottom. Then the film was hydrated with 1 mL of PLGA nanoparticles (5 mg/mL) and was subsequently extruded 11 times through a 200 nm polycarbonate membrane using Avanti Mini-Extruder, forming the Lipo-NPs.


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Characterization of the nanoparticles. To examine the nanoparticle morphology, 10 µL of sample solution was dropped onto carbon-coated copper grids and was deposited for 3 min. After the residual fluid was absorbed using filter paper, samples on the grids were negatively stained with 1% (v/v) uranyl acetate for 2 min and air dried for analysis using a transmission electron microscope (TEM, HT7700, HITACHI, Japan). The hydrodynamic size distribution, zeta potential and polydispersity index (PDI) of nanoparticles were assessed using a Zetasizer Nano ZS dynamic light scattering (DLS) instrument (Malvern, UK). Analysis of MSC membrane proteins. The protein concentrations of MSC cell lysate, plasma membrane and PM-NP were quantified using BCA assay. These samples with equivalent amount of total proteins were mixed with loading buffer, and were separated in a 10% Bis-Tris stain-free mini-gel using Bio-Rad electrophoresis system (USA). After electrophoresis, the protein pattern in the gel was visualized using Bio-Rad ChemiDox Touch Imaging System (USA). The expression of membrane markers C-X-C chemokine receptor type 4 (CXCR4) and Na+-K+ ATPase in these samples were identified using western blot. In brief, proteins on the gel were transferred to a PVDF membrane. Then the PVDF membrane was blocked with 5% non-fat milk for 1 h, incubated with primary antibodies overnight. After washing in Tris-buffered saline containing 0.02% Tween-20 (TBST) for three times, the membrane was incubated with secondary antibodies for 1 h, followed by washing with TBST for three times. The PVDF membrane was incubated with ECL substrate followed by exposure under Bio-Rad ChemiDox Touch Imaging System. Drug encapsulation efficiency and in vitro release. The drug encapsulation efficiency and drug loading content were calculated using the following formulas:


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Encapsulation efficiency (%) = Mass of drug loaded by nanoparticles/Initial mass of drug × 100% Loading content (%) = Mass of drug loaded by nanoparticles/(Mass of drug loaded by nanoparticles + mass of nanoparticles) × 100% To measure the drug release profile, 1 mL of PM-NP-Dox solution (10 mg/mL in PBS) was dialyzed against 50 mL of PBS at pH 7.4 or pH 4.5 using a dialysis bag (molecular weight, 2000 Da) at 37 oC. At different time intervals, 1 mL of the dialysis buffer was used to detect the amount of Dox. For above experiments, the amount of Dox was determined by detecting the absorbance value at the wavelength of 485 nm using a UV-Vis spectrophotometer (AOE Instruments, Shanghai, China). Cellular uptake of the nanoparticles. MHCC97H cells were incubated with PM-NPRhodamine B for 1 h or 4 h. Then the cells were washed twice with PBS and cell pellets were collected by centrifugation at 200 g. The fluorescence signals in cells were analyzed using flow cytometry (BD Accuri C6, USA). To analyze the cellular uptake behavior of plasma membrane and PLGA nanoparticle core in PM-NPs, dual-labeled PM-NPs were prepared by loading PLGA nanoparticles with Rhodamine B and labeling plasma membrane with PKH67. MHCC97H cells were seeded onto 35 mm glass-bottom confocal dishes at a density of 1×105 cells/dish and were cultured for 24 h. Then the cells were incubated with dual-labeled PM-NPs for 1 h. Fluorescence signals were examined using confocal microscopy (Zeiss 710, Germany). To further investigate the cellular uptake process and intracellular colocalization of PM-NPs, the cells were incubated with PM-NPs loaded with Rhodamine B for 1 h and 4 h. Lysosensor green DND was added into the culture medium for labeling the endo-lysosomes. After washed three times with PBS, the cells were examined using confocal microscopy.


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Cytotoxicity. Cell viability was assessed based on CCK-8 assay. MHCC97H cells were seeded into 96-well plates at a density of 5 × 103 cells/well and were cultured overnight. Then the cells were treated with Dox, NP-Dox, Lipo-NP-Dox or PM-NP-Dox with the equivalent Dox concentration of 0, 0.2, 0.5, 1, 2, or 5 µM. After 24 h, the cells were washed twice with PBS and incubated with CCK-8 solution for 1 h. The supernatant was transferred to a new 96-well plate to exclude the interference of intracellular Dox on CCK-8 absorbance. The absorbance at 450 nm was detected using a microplate reader (Perkin Elmer, UK). The cytotoxicity of the nanocarrier itself (PM-NP) was also evaluated using CCK-8 assay. MHCC97H cells and HUVECs were each seeded onto 96-well plates at a density of 5 × 103 cells/well and cultured overnight. The cells were treated with PM-NP at the concentration of 0, 2.9, 7.25, 14.5, 29, and 72.5 µg/mL (equal to PLGA concentration). After 24 h, the cells were incubated with CCK-8 solution for 1 h. The absorbance at 450 nm was detected. Animals and tumor model. All animal studies were approved by the Institutional Animal Care and Use Committee of National Center for Nanoscience and Technology. Female BALB/c nude mice (6-8 weeks) were purchased from Vital River Animal Laboratories (Beijing, China). To establish a xenograft liver cancer model, about 1 × 106 MHCC97H cells were suspended in 100 µL of PBS/matrigel mixture, and were inoculated subcutaneously at the back of each nude mouse. In vivo and ex vivo imaging of PM-NPs. PM-NPs and Lipo-NPs were both loaded with ICG fluorescence molecules and were then injected to nude mice bearing subcutaneous MHCC97H liver tumors through the tail vein. After different time intervals (4, 8 or 20 h), fluorescence signals were examined using an in vivo imaging system (IVIS Spectrum CT, Perkin Elmer, UK). Twenty hours after injection, the mice were euthanized and tumors and major organs were


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separated for ex vivo imaging of the nanoparticles. The fluorescence intensity of region of interest (ROI) was analyzed by the imaging system software. Antitumor efficacy. When the average volume of tumor xenografts reached about 100 mm3, the tumor-bearing mice were divided into 4 groups (5 mice/group) and treated with saline, NP-Dox, Lipo-NP-Dox or PM-NP-Dox (equivalent concentration of Dox, 4 mg/kg) every three days through tail vein injection. Tumor size and body weight of mice were measured every three days. Tumor volume was calculated using the formula: Volume = (length×width2)/2. Immunohistochemistry and immunofluorescence. After treatments, tumors and major organs including hearts, livers, spleens, lungs and kidneys were harvested and fixed in 4% paraformaldehyde solution. Then, all the tissues were dehydrated with ethanol at serial concentrations, transparentized with xylene and embedded in paraffin. Paraffin sections (7 µm) were dewaxed, rehydrated and incubated with 0.3% H2O2 in methanol. For hematoxylin and eosin (H&E) staining, tissue sections were stained with Mayer’s hematoxylin for 3 min and eosin Y for 2 min, and observed using light microscopy (Leica, Germany). For immunofluorescence, tumor sections were incubated with protease K (10 µg/mL, pH 7.4) at 37 oC for 15 min. After rinsed twice with PBS, the slices were incubated with FITC-labeled TUNEL reaction reagent for 1 h at 37 oC in a humidified atmosphere in the dark. After washed five times with PBS, the slices were stained with DAPI and examined using confocal microscopy.

RESULTS AND DISCUSSION Characterization

of

umbilical

cord-derived

MSC

plasma

membrane-modified

nanoparticles. Considering the drug loading capacity and biocompatibility, we chose an FDA-


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approved co-polymer PLGA for construction of the nanoparticle core through a well-studied double-emulsion method. Umbilical cord-derived MSC plasma membrane was isolated for coating the PLGA nanoparticles, forming a hybrid nanoparticle PM-NP. TEM examination revealed that PLGA nanoparticles exhibited a uniform and spherical nanostructure and the plasma membrane showed a typical lipid bilayer (Figure 1A). The hybrid PM-NP was characterized by a spherical core-shell structure with lipid bilayer on the surface (Figure 1A). DLS analysis showed that PLGA nanoparticles had a hydrodynamic size of 80.9 ± 0.6 nm and a ζ-potential of -50.8 ± 2.1 mV (Figure 1B). After modification with the plasma membrane, the hydrodynamic diameter of the nanoparticles reached 103.6 ± 2.9 nm and the surface charge varied to -30.3 ± 2.6 mV, which was close to that of the plasma membrane vesicles (Figure 1B), suggesting that PLGA nanoparticles were successfully coated with MSC plasma membranes. The negative surface charge of PM-NPs would protect the nanoparticles from non-specific absorption of proteins, ensuring the stability of PM-NPs in blood circulation.37, 38 To introduce a control of PM-NPs, we constructed a liposome-coated PLGA nanoparticle (Lipo-NP) that exhibited similar morphology, size, and surface charge compared with PM-NP (Figure 1A and 1B). To determine the optimal amount of surface coated plasma membrane, we prepared PMNPs with PLGA and plasma membrane at various mass ratios. The ζ-potential gradually increased as the amount of plasma membrane increased (Table 1). When the mass ratio of plasma membrane protein and PLGA nanoparticles was 1: 5, the surface charge achieved -30.6 mV that was very close to the charge of plasma membrane vesicles (Table 1), indicating that the coating of plasma membrane was saturated at this ratio. Since proteins on MSC plasma membrane especially chemokine receptors are essential for the tumor-targeting feature of MSCs,


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we compared the protein expression pattern of pure MSC plasma membranes and PM-NPs using SDS-PAGE and western blot assays. SDS-PAGE gel analysis revealed that the protein profiles of PM-NPs and purified plasma membranes were similar, while they were distinct from that of whole cell lysates (Figure 1C). Meanwhile, the expression levels of the well-known membrane protein Na+/K+ ATPase and the chemokine receptor CXCR4 were identical between PM-NPs and MSC plasma membranes (Figure 1D). The closely matched protein profiles of PM-NPs and pure plasma membranes suggested that membrane proteins were not lost or degraded during the coating process and further verified that the nanoparticles were modified with MSC plasma membranes. Characterization of drug-loaded PM-NPs. The well-defined PM-NPs were utilized to encapsulate a hydrophilic chemotherapeutic drug doxorubicin (Dox, denoted as PM-NP-Dox) for xenograft liver tumor treatment. To optimize the drug encapsulation efficiency, PLGA and Dox at different mass ratios were used for construction of PM-NP-Dox (Table 2). At the proportion of 25: 3 (mass ratio of PLGA: Dox), PM-NP-Dox showed an appropriate particle size (104.2 nm) and a low PDI value (0.168), and most importantly, relatively high encapsulation efficiency (21.8%) and drug loading content (2.3%). Thus, this mass ratio was chosen for construction of PM-NP-Dox in further studies. TEM and DLS analysis revealed that PM-NP-Dox was core-shell structured spherical nanoparticle with a hydrodynamic size of 105 nm and a ζ-potential of -27.2 ± 9.58 mV (Figure 2A and 2B). The stability of PM-NP-Dox and Lipo-NP-Dox in PBS buffer (pH 7.4) was examined by measuring the size and ζ-potential at various time points (Figure 2C). Only slight changes of the two parameters were observed after incubation for 7 days, indicating that the plasma membrane or liposome modified nanoparticles were stable in physiological environment. Taken together, these results demonstrate that MSC plasma membrane-modified


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PLGA nanoparticles were successfully constructed with desirable features for in vivo drug delivery. Cellular uptake of PM-NP-Dox in vitro. Since PM-NP-Dox was hypothesized to enter target cells, we next examined the cellular uptake profile of PM-NPs. Dual-labeled PM-NPs were prepared by loading PLGA nanoparticles with Rhodamine B and labeling plasma membrane with the green lipophilic fluorescent dye PKH67, followed by incubation with MHCC97H cells for 1 h. Confocal microscopy observation revealed that PM-NPs could enter cells, and the nanoparticle core and plasma membrane were mostly co-localized (Figure 3A), suggesting that PM-NPs remained intact during the cellular uptake process. To further investigate the effect of plasma membrane functionalization on cellular uptake of nanoparticles, we incubated MHCC97H cells with Rhodamine B-labeled PLGA nanoparticles, Lipo-NPs, or PM-NPs for 1 h or 4 h, and analyzed the intracellular amount of the nanoparticles by flow cytometry (Figure 3B). The cellular uptake efficiency of PM-NPs was higher than that of bare PLGA nanoparticles or LipoNPs, and the differences became more obvious in accordance with incubation time. After incubation for 4 h, the amount of the internalized PM-NPs was about 2 times of PLGA nanoparticles and 3 times of Lipo-NPs, indicating that surface coating of plasma membrane can aid nanoparticle cellular uptake. The variation in cellular uptake efficiency of PM-NPs and LipoNPs likely resulted from the enhanced interaction between PM-NPs and the cell membrane. We also evaluated the subcellular localization of PM-NPs by incubation of MHCC97H cells with Rhodamine B-labeled PM-NPs and subsequent staining of endo-lysosomes. Confocal microscopy analysis revealed that internalized PM-NPs gradually accumulated in the acidic organelles where they can release cargos effectively (Figure 3C). These results demonstrate that PM-NP-Dox can be efficiently taken up by target cells and translocate to acidic organelles.


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Drug release profile and antitumor effects of PM-NP-Dox in vitro. Then the drug release profile of PM-NP-Dox was evaluated in PBS buffer at pH 7.4 and pH 4.5 (Figure 4A), the conditions mimicking the physiological environment and endo-lysosomal compartment interior environment.39 At pH 7.4, the accumulated drug release achieved over 76% in Dox-loaded PLGA nanoparticles (NP-Dox) and was 47% in Dox-loaded Lipo-NPs (Lipo-NP-Dox) after 52 h. In contrast, less than 35% of Dox was released from PM-NP-Dox under the same condition, indicating that surface modification of plasma membrane can protect the drugs from burst release in blood circulation. At pH 4.5, the drug release behaviors of PM-NP-Dox and Lipo-NP-Dox were similar and over 60% of Dox was released within 5 h, suggesting that the acidic environment was conducive to drug release. We next examined the cytotoxicity of PLGA nanoparticles, Lipo-NPs, and PM-NPs. CCK-8 analysis showed that these nanocarriers had no obvious cytotoxic effect even at the concentration of 72.5 µg/mL (Figure S1). To investigate the antitumor activity of PM-NP-Dox in vitro, MHCC97H cells were treated with free Dox, NP-Dox, Lipo-NP-Dox, or PM-NP-Dox for 24 h and cell viability was evaluated using CCK-8 assay. Approximately 70% cancer cells were killed after treatment with PM-NP-Dox at the concentration of 2 µM (Figure 4B). The IC50 of Dox, NP-Dox, Lipo-NP-Dox, and PM-NP-Dox was 1.88 µM, 1.40 µM, 1.41 µM, and 1.09 µM, respectively (Figure 4C). The tumor-killing capacity of these nanoformulations correlated with the cellular uptake efficiency of relevant nanocarriers. Moreover, the apoptosis-inducing effect of above treatments was further evaluated by examining the expression of cleaved caspase-3 and total caspase-3 in cells. Figure 4D showed that the level of cleaved caspase-3 was higher in PM-NP-Dox treated cells than those in other groups. These results demonstrate that PM-NP-Dox can efficiently release drugs in acidic environment and thereby kill tumor cells.


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Stability and tumor targeting of PM-NPs in vivo. To evaluate the stability of PM-NPs in blood circulation, we prepared PM-NPs encapsulating Cy7 fluorescence molecules (named PM-NPCy7) and injected free Cy7 or PM-NP-Cy7 to mice through the tail vein. After different time intervals, blood was collected and the fluorescence intensities were examined using in vivo imaging system to indicate the pharmacokinetics of PM-NPs (Figure S2A). Based on this method, the circulatory half-life of PM-NP-Cy7 and free Cy7 was about 2 h and 0.6 h, respectively (Figure S2B). Since surface functionalization of plasma membrane was considered to endow PLGA nanoparticles with intrinsic tumor-targeting features, tumor accumulation and the biodistribution of PM-NPs were therefore examined. PM-NPs and Lipo-NPs were encapsulated with a near-infrared fluorescent probe ICG, and were intravenously injected to BALB/c nude mice bearing MHCC97H liver tumor xenografts. After various time intervals, the fluorescent signals were analyzed using an in vivo imaging system. As shown in Figure 5A, fluorescence was observed across the body in both groups 4 h or 8 h after injection. After 20 h, most fluorescent signals were distributed at the tumor sites due to the metabolism and tumor accumulation of the PM-NPs. Intriguingly, the fluorescence intensity of tumor-localized PM-NPs was about 2 times higher than that of Lipo-NPs after 20 h (Figure 5B), suggesting that PM-NPs exhibited active tumor-targeting traits besides the passive targeting effect of nanoparticles. To quantitatively evaluate the tumor-targeting efficiency of PM-NPs, tumors and major organs including hearts, livers, spleens, lungs, and kidneys were used for ex vivo imaging. Comparable fluorescence signals of PM-NPs and Lipo-NPs were observed in hearts, spleens, lungs, and kidneys (Figure 5C and 5D). However, the fluorescence intensity of PM-NPs in tumors was about 2 times higher than that of Lipo-NPs (Figure 5C and 5D). To further demonstrate the role of MSC plasma membrane in the tumor targeting of PM-NPs, we prepared PLGA nanoparticles


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coated with human red blood cell membranes (named RBC PM-NP) and utilized it as a control when evaluating the biodistribution of PM-NPs. RBC PM-NP also exhibited spherical nanoparticle morphology (Figure S3A), and showed a hydrodynamic diameter of 105.1 nm (Figure S3B) and a zeta potential of -26.7 mV (Figure S3C). PM-NPs and RBC PM-NPs were labeled with Cy7 fluorescence molecules and were injected to tumor-bearing nude mice through the tail vein. Twenty hours after injection, tumors and main organs were separated for ex vivo imaging. We observed that the fluorescence intensity of PM-NPs was about 2 times higher than that of RBC PM-NPs within tumors (Figure S3D and S3E), further verifying the contribution of MSC plasma membrane to the active tumor targeting of nanoparticles. The tumor-targeting effect of PM-NPs was likely attributed to the expression of many chemokine receptors such as plateletderived growth factor receptor (PDGFR) and CXCR4 that can bind to corresponding chemokines in tumor tissues22, 40, 41. Nevertheless, the detailed mechanism requires extensive investigation. Meanwhile, lower fluorescence levels of PM-NPs were detected in the livers compared with that of Lipo-NPs (Figure 5C and 5D), indicating that MSC plasma membrane can protect nanoparticles from elimination by the reticuloendothelial system in the liver. The underlying mechanism possibly lay in the constitutive expression of CD47 molecules on the surface of MSC plasma membrane.31 CD47 has been reported to bind with signal-regulatory protein α on macrophages and Kupffer cells, and deliver the “do not eat me” signal.31, 42, 43 Therefore, PM-NP with CD47 molecules on the surface is likely to escape from the phagocytosis of Kupffer cells in the liver. These data demonstrate that surface modification of MSC plasma membrane can increase the tumor targeting and accumulation of nanoparticles and decrease the possibility of non-specific elimination.


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Antitumor efficacy and biosafety of PM-NP-Dox in vivo. Based on the superiority of PM-NPs on tumor accumulation, we then investigated the antitumor effect of PM-NP-Dox in BALB/c nude mice with subcutaneous MHCC97H liver tumors. When the average tumor volume was approximately 100 mm3, mice were divided into four groups and were injected with saline, NPDox, Lipo-NP-Dox, or PM-NP-Dox (equal to 4 mg/kg of Dox) every 3 days for 5 times through the tail vein. From the tumor growth curve in Figure 6A and tumor images in Figure 6B, both NP-Dox and Lipo-NP-Dox showed certain tumor inhibition activity. In contrast, PM-NP-Dox dramatically inhibited fast tumor growth. When treatments were terminated, the tumor inhibition rates of NP-Dox, Lipo-NP-Dox, and PM-NP-Dox were 34.4%, 32.8% and 78.2%, respectively. Measurement of tumor weights further verified the evident antitumor effect of PM-NP-Dox (Figure 6C). The highest antitumor efficacy of PM-NP-Dox was attributable to the tumortargeting capacity of PM-NPs. The apoptosis status of tumor tissue was then analyzed by H&E staining and TUNEL staining of tumor tissue sections. As shown in Figure 6D, massive karyolysis and apoptosis areas were observed in tumor sections of PM-NP-Dox treated group. TUNEL staining also revealed the highest DNA damage ratio in PM-NP-Dox treated tumors (Figure 6E and 6F). The tumor apoptosis status was consistent with the antitumor efficacy of various treatments. The aforementioned data demonstrate that PM-NP-Dox hold great potential in shrinking tumor fast growth by efficient tumor-targeted drug delivery. The accumulation of PM-NPs in livers and kidneys prompted us to further investigate the possible adverse effects of PM-NP-Dox. The body weights of the above treated mice were monitored and no abnormality was observed in each treatment group (Figure 7A). Moreover, H&E staining of main organ sections (heart, liver, spleen, lung, and kidney) showed no obvious


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changes in morphology (Figure 7B). These preliminary results proved the safety profile of PMNP-Dox in vivo.

CONCLUSIONS In summary, our present work reported a potent drug delivery system consisting of PLGA nanoparticle core and surface-coated plasma membrane from umbilical cord-derived MSCs. The nanoparticles could be taken up by tumor cells and accumulated in acidic organelles for drug release, thereby inducing tumor cell death. Moreover, the nanocarriers achieved tumor-targeted delivery of chemotherapeutic drugs, resulting in effective tumor suppression. This study for the first time utilized umbilical cord-derived MSC plasma membrane for nanoparticle functionalization, and provided a promising vehicle for tumor-targeted drug delivery with potential for personalized cancer targeting and treatment.


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Figure 1. Characterization of MSC plasma membrane functionalized PLGA nanoparticles (PM-NP). (A) TEM images of PLGA nanoparticles (PLGA NP), plasma membrane (PM) vesicles, plasma membrane-coated PLGA nanoparticles (PM-NP), and liposome-coated PLGA nanoparticles (Lipo-NP). Scale bar, 100 nm. Magnified images in PM-NP and Lipo-NP are inserted. Scale bar, 40 nm. The arrows indicated the typical lipid bilayers. (B) The average hydrodynamic diameters and surface charges of PLGA NP, PM vesicle, PM-NP, liposome, and Lipo-NP were examined using DLS. All the data are presented as mean ± s.d. (n = 3) (C) The expression profile of proteins in PM-NP, PM, and whole cell lysates of MSCs were analyzed using SDS-PAGE. (D) The expression levels of two membrane proteins (Na+-K+ ATPase and CXCR4) in PM-NP and PM were evaluated using western blot.


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Figure 2. Characterization of PM-NP-Dox. (A) TEM image of PM-NP-Dox revealed a spherical core-shell nanostructure. Scale bar, 100 nm. (B) DLS analysis of the PM-NP-Dox size revealed an average diameter of 104.2 nm (PDI = 0.168). (C) The stability of PM-NP-Dox was evaluated by monitoring variation in size and surface charge after standing at room temperature for different time intervals. All the data are presented as mean ± s.d. (n = 3).


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Figure 3. Cellular uptake of PM-NPs. (A) Co-localization of PM and PLGA NPs was observed under confocal microscopy after dual-labeled PM-NPs were incubated with MHCC97H cells for 1 h. For PM-NP, the PM component was labeled with PKH67, and PLGA NPs was loaded with Rhodamine B. Nucleus was stained by Hoechst 33342. Scale bar, 20 µm. (B) Quantification of internalized PLGA-NPs, Lipo-NPs, and PM-NPs using flow cytometry after these nanoparticles


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were incubated with cells for 1 h or 4 h. All the data are presented as mean ± s.d. (n = 3) and analyzed with two-way ANOVA (**, P < 0.01). (C) Co-localization of PM-NPs and endolysosomes were analyzed using confocal microscopy. PM-NPs were loaded with Rhodamine B and endo-lysosomes were labeled using FITC-lysosensor. Scale bar, 20 µm.


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Figure 4. The drug release profile and antitumor effects of PM-NP-Dox in vitro. (A) The drug release profiles of Dox-loaded PLGA nanoparticles (NP-Dox), Dox-loaded Lipo-NP (LipoNP-Dox), and Dox-loaded PM-NP (PM-NP-Dox) at pH 7.4 or pH 4.5. All the data are presented as mean ± s.d. (n = 3). (B) The tumor cell killing effects of PM-NP-Dox was revealed by CCK-8


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analysis. Cells were treated with Dox, NP-Dox, Lipo-NP-Dox, or PM-NP-Dox at different concentrations for 24 h. Data are presented as mean ± s.d. (n = 6) and analyzed with two-way ANOVA (*, P < 0.05; **, P < 0.01). (C) The IC50 of formulations in panel B. Data are presented as mean ± s.d. (n = 6) and analyzed with one-way ANOVA (**, P < 0.01; ***, P < 0.001). (D) The expression of cleaved caspase 3 and caspase 3 in MHCC97H cells that were treated with NP-Dox, Lipo-NP-Dox, or PM-NP-Dox (equal to 2 µM Dox) was analyzed using western blot. Positive control sample was prepared by incubating MHCC97H cells at 55 oC for 20 min.


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Figure 5. Biodistribution of PM-NP in tumor-bearing mice. (A) In vivo imaging of PM-NP and Lipo-NP in mice bearing MHCC97H liver tumor xenograft. PM-NP and Lipo-NP were loaded with ICG for near-infrared imaging. Excitation wavelength, 745 nm; emission wavelength, 820 nm. (B) Quantification of the tumor regional fluorescence signals in panel A. Data are presented as mean ± s.d. (n = 3) and analyzed with two-tailed t-test (*, P < 0.05; **, P < 0.01). (C) Ex vivo imaging of PM-NP and Lipo-NP in tumors and major organs. (D) Quantification of the fluorescence signals in panel C revealed the relative amount of PM-NP and Lipo-NP in tumors and various organs. Data are presented as mean ± s.d. (n = 3) and analyzed with two-tailed t-test (*, P < 0.05; **, P < 0.01).


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Figure 6. Antitumor efficacy of PM-NP-Dox in vivo. (A) The growth curves of liver tumors treated with saline, NP-Dox, Lipo-NP-Dox, or PM-NP-Dox. Data are presented as mean ± s.d. (n = 5). The tumor volumes in different treatment groups on day 15 were analyzed with one-way ANOVA (*, P < 0.05; **, P < 0.01). (B) Images of tumors were captured after tumor-bearing mice received four treatments for 15 days. (C) Tumor weights were measured when the


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treatments were terminated. All the data are presented as mean ± s.d. (n = 5) and analyzed with two-tailed t-test (*, P < 0.05; **, P < 0.01). (D) H&E staining of tumor sections in panel B. Scale bar, 50 µm. (E) The apoptosis status of tumor tissues in panel B was analyzed by TUNEL staining. Blue: nuclei, green: TUNEL positive area. Scale bar, 100 µm. (F) The apoptosis degree was quantified by statistical analysis of TUNEL positive areas in panel E. All the data are presented as mean ± s.d. (n = 5) and analyzed with one-way ANOVA (**, P < 0.01).


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Figure 7. The biosafety of PM-NP-Dox. (A) The body weights of tumor-bearing mice were monitored during treatment period (15 days). Data are presented as mean ± s.d. (n = 5). (B) H&E staining of major organs from tumor-bearing mice when the treatments were terminated. Scale bar, 50 µm.


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Table 1. The ζ-potential values of PM, PLGA NP and PM-NP-Dox PM-NP (PM protein: NPs, w/w) PM Zeta potential (mV)

PLGA NP

-30.3±2.6

-50.3±1.9

1: 20

1: 10

1: 5

-40.0±1.0

-35.6±2.4

-30.6±3.8

Table 2. Drug loading efficiency and loading content of PM-NP-Dox at different mass ratio of PLGA and Dox PLGA input (mg) 10

Dox input (mg) 0.4

Drug encapsulation efficiency 38.0%

Drug loading content 1.5%

10

0.8

26.9%

10

1.0

10 10

Size (nm)

PDI

80.59

0.092

2.0%

86.34

0.106

24.3%

2.2%

85.49

0.121

1.2

21.8%

2.3%

104.20

0.168

1.4

19.3%

2.4%

134.40

0.429


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AUTHOR INFORMATION Corresponding Authors *Guangjun Nie, Email: [email protected]; *Yanping Ding, Email: [email protected]. *Wen-Fei Dong, Email: [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by the grants from the National Basic Research Plan of China (2018YFA0208900), the National Natural Science Foundation of China (31671023, 31700864, 31571021, 11621505, 81771982), Beijing Natural Science Foundation (7174333), the Key Research Program of Frontier Sciences, CAS (QYZDJ-SSW-SLH022), the Key Research Program of CAS (KFZD-SW-210), and the National Key R&D Program of China (2017YFF0108600).

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Surface Functionalization of Polymeric Nanoparticles with Umbilical

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