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Targeted lipid nanoemulsions encapsulating epigenetic drugs exhibit selective cytotoxicity on CDH1-/FOXM1+ triple negative breast cancer cells Bumjun Kim, Caroline D. Pena, and Debra T. Auguste Mol. Pharmaceutics, Just Accepted Manuscript • DOI: 10.1021/acs.molpharmaceut.8b01065 • Publication Date (Web): 18 Mar 2019 Downloaded from http://pubs.acs.org on March 19, 2019

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Molecular Pharmaceutics

Targeted lipid nanoemulsions encapsulating epigenetic drugs exhibit selective cytotoxicity on CDH1-/FOXM1+ triple negative breast cancer cells Bumjun Kima,b, Caroline D. Penaa, Debra T. Augustea,b, 1 a Department

of Biomedical Engineering, The City College of New York, 160 Convent Avenue, New York, NY, 10031, United States

b Current: Department of Chemical Engineering, Northeastern University, 360 Huntington Avenue, Boston, MA 02115, United States 1 Corresponding author: Professor. Debra T. Auguste, Tel: 617-373-6243, Email: [email protected]

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Abstract The plasticity of cancer epigenetics makes them plausible candidates for therapeutic intervention. We took advantage of elevated expression of lysophosphatidic acid receptor 1 (LPAR1) in triple negative breast cancer (TNBC) tissues to target Decitabine (DAC) and Panobinostat (PAN) to breast cancer cells. DAC and PAN were shown to reverse abnormal methylation of DNA and altered chromatin structure, respectively, leading to increased expression of tumor suppressor genes and decreased expression of oncogenes. Although DAC and PAN have therapeutic benefits, they are limited by chemical instability and systemic toxicity. Herein, we present LPAR1-targeted, lipid nanoemulsions (LNEs) encapsulating both DAC and PAN. Our results demonstrated that the cell uptake and in vivo biodistribution of LNEs was dependent on LPAR1 expression in TNBCs. DAC/PAN-LNEs were effective in inhibiting the growth of mesenchymal breast cancer cells by restoring CDH1/E-cadherin and suppressing FOXM1 expression. Epithelial breast cancer cells that inherently express low FOXM1 and high CDH1 were unaffected by DAC/PAN-LNEs. Overall, we successfully designed LPAR1targeted LNEs that selectively act on CDH1(low)/FOXM1(high) TNBC cell lines.

Keywords: Lipid nanoemulsions, LPA, Triple negative breast cancers, Epigenetic drug, Panobinostat, Decitabine, CDH1, FOXM1


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Introduction Breast cancer is a heterogeneous disease that includes a variety of subtypes with unique morphologies and clinical behaviors[1, 2]. Some basal-like cancers, called triple negative breast cancer (TNBC), lack three distinctive biomarkers - human epidermal growth factor receptor 2 (HER2), estrogen receptor (ER), and progesterone receptor (PR)[3]. TNBCs constitute 10 % to 20 % of breast cancer cases, and show more aggressiveness, metastasis, recurrence and poorer prognosis than non-TNBCs[4, 5]. Conventional chemotherapeutics lack specificity to the diseased cells, resulting in severe off-target effects[6, 7]. Thus, the absence of molecular biomarkers and toxicity of chemotherapeutics suggests an urgent and unmet need for therapeutic alternatives for treating TNBCs[8]. Cancers present aberrant reprogramming of epigenetic machinery such as DNA hypermethylation and histone deacetylation, contributing to cancer initiation and progression[9, 10]. DNA methylation is catalyzed by DNA methyltransferases (DNMTs); an increase in DNMTs results in hypermethylation of promoter CpG islands, leading to stable gene silencing[11]. Tumor suppressor genes are inactivated by promotor CpG island hypermethylation[12, 13]. These genes can be activated by DNMT inhibitors (DNMTis) such as 5-Aza-2’ –deoxycytidine (DAC)[14]. DAC is incorporated into DNA and covalently traps DNMTs on DNA, resulting in the demethylation of DNA[15]. Global histone deacetylation, mediated by histone deacetylases (HDACs), is another hallmark of abnormal cancer epigenetics, which leads to gene repression[16]. Increased expression of HDACs is measured in different cancers; treatment with HDAC inhibitors (HDACis) suppress the activity of HDACs, leading to the upregulation and downregulation of tumor suppressor genes and oncogenes, respectively. HDACi treatment demonstrated success


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in preclinical and clinical models of multiple myeloma, non-small cell lung cancer, prostate cancer, cervical cancer, and ovarian cancer [17-19]. Panobinostat (PAN), among the other HDAC inhibitors, is shown to selectively inhibit the growth of TNBCs at nanomolar concentrations[20]. Evidence has suggested that the combined treatment of DAC and PAN can synergistically reverse the aberrant epigenetics, resulting in enhanced therapeutic efficiency in a wide range of cancers[21-24]. Co-delivery of DAC and PAN is difficult due to their different chemistries, PAN is hydrophobic and DAC is hydrophilic. Rapid degradation of DAC in aqueous solution and its high clearance rate in vivo limits its application to treat solid tumors[25, 26]. The severe systemic toxicity of PAN, such as thrombocytopenia, anemia, and neutropenia, limits its use[27, 28]. We hypothesized that DAC and PAN formulated into a targeted drug delivery vehicle may effectively increase tumor suppressor genes and decrease oncogenes, protect DAC from degradation, and limit the off-target toxicity of PAN. Lipid nanoemulsions (LNEs) LNEs are nanoscale oil-in-water emulsions, with diameters in the range of 20-200nm[29]. Due to their ease of fabrication, biocompatibility, and long term stability[30], LNEs are considered useful vehicles for delivery of hydrophobic molecules. Soybean oil is widely used to create LNEs; however, soybean oil contains far more omega-6 polyunsaturated fatty acids (PUFAs) than omega3 PUFAs[31]. Excessive omega-6 PUFA consumption is associated with many chronic diseases prevalent in modern society including cancer, whereas the increased intake of omega-3 PUFA and decreased intake of omega-6 PUFA exhibited beneficial effects on patients with these diseases[32]. In contrast, cod liver oil (CLO) is a good source of omega-3 PUFAs. More than 20 % of PUFAs in CLO are omega-3 PUFAs, whereas only 4 %


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are omega-6 PUFAs[33]. Omega-3 PUFA intake is associated with lower risk of cancer and death in cancer patients[34-36]. LNEs are advantageous as their size, below 200 nm, selectively targets tumors through the enhanced permeability and retention (EPR) and provides improved stability of drugs by limiting enzymatic and hydrolytic degradation[37]. This may lead to an increase in effective drug concentration in tumor tissues[38]. The ability of LNEs to target tumors may be further improved by the addition of tumor-targeting ligands[39-41]. Increased synthesis of lysophosphatidic acid (LPA) and its receptors (LPAR1-3) are implicated in breast cancers[42]. LPA/LPAR1 has been linked to breast cancer metastasis both in vitro and in vivo[43-45]. Higher mRNA expression of LPAR1 in primary tumor of breast cancer patients significantly correlated with lymph node invasion[46]. The LPA/LPAR1-3 axis is reported to affect angiogenesis, metastasis, and proliferation[42, 47]. Overexpression of LPAR1-3 in transgenic mice resulted in mammary carcinoma[48]. Due to the emerging role of the LPA/LPAR1-3 axis in cancer progression, blocking LPA/LPAR1-3 signaling has shown potential as an anti-cancer therapy[49]. G2A, a receptor activated by lysophosphatidylcholine (LPC), is also overexpressed in human breast tumor samples[50]. G2A is a member of the G protein-coupled receptors; higher expression of G2A is clinically associated with lower metastasis free survival in breast cancer[51]. LPC is the precursor of LPA; LPC is also reported to antagonize pH-dependent activation of G2A[52]. However, the study of LPAR1-3 or G2A as a molecular target for drug delivery has yet to be examined. In this study, we measured the impact of dual-delivery of DAC and PAN, using LPAR1-targeted LNEs comprised of different ratios of LPA to LPC. Cellular uptake and biodistribution of LNEs in TNBCs were investigated. The mechanism of epigenetic drug


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encapsulating LNEs was verified by lowering expression of an oncogene and increasing expression of a tumor suppressor gene. Our findings demonstrated that epigenetic drugs delivered via LPAR1-targeted LNEs may be an alternative therapeutic option for TNBCs.


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Materials and Method Chemicals 1-oleoyl-2-hydroxy-sn-glycero-3-phosphate, 1-oleoyl-2-hydroxy-sn-glycero-3phosphocholine, and 1, 2-distearoyl-sn-glycero-3-phosphoethanolamine-N[carboxy(polyethylene glycol)-2000 were all obtained from Avanti Polar Lipids (Alabaster, AL). 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindotricarbocyanine iodide (DiR) was acquired from Biotium (Fremont, CA). Cod liver oil was purchased from MP Biomedicals (Irvine, CA). Decitabine and Panobinostat were acquired from Selleck Chemicals (Huston, TX). Rhodamine 123 was purchased from Enzo Life Science (Farmingdale, NY). Nanosep Centrifugal Device Omega was obtained from Pall (Port Washington, NY). FITCconjugated rabbit polyclonal anti-EDG2 antibody (LPAR1 antibody) and FITC-conjugated rabbit polyclonal anti-G2A antibody were acquired from Bioss (Woburn, MA). Primer pairs of B2M, CDH1, and FOXM1 were obtained from Integrated DNA Technologies (Coralville, IA). Isotype Control IgG (tagged with Alexa Fluor®647) and mouse anti-human CD324 (tagged with Alexa Fluor®647) were both purchased from Biolegend (San Diego, CA). Rabbit FOXM1 monoclonal antibody (D12D5) was purchased from Cell Signaling Technology (Danvers, MA). Mouse HPRT1 monoclonal antibody (F-1) and Mouse IgG kappa binding protein (m-IgGĸ BP, sc-516102) were obtained from Santa Cruz Biotechnology (Dallas, TX). Amersham ECL Rabbit IgG antibody (NA934V) was acquired from GE Healthcare Life Sciences (Marlborough, MA). MEGM Bullet kit and media were purchased from Lonza (Allendale, NJ). Dulbecco’s modified Eagle medium (DMEM) and DMEM/F12 was acquired from GE Healthcare Life Sciences (Piscataway, NJ). Dulbecco’s phosphate buffered saline (PBS), Leibovitz's L-15 Medium, Roswell Park Memorial Institute (RPMI)-1640 Medium, and 0.25 % trypsin/2.21 mM ethylenediaminetetraacetic


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acid (EDTA) solution were obtained from ATCC (Manassas, VA). Horse serum (HS), fetal bovine serum (FBS), and penicillin-streptomycin (PS) were obtained from Thermo Fisher Scientific (Waltham, MA). Epidermal growth factor (EGF) was purchased from Peprotech (Rocky Hill, NJ). Hydrocortisone, cholera toxin, insulin, resazurin sodium salt, and dimethyl sulfoxide (DMSO) were purchased from Sigma-Aldrich (St. Louis, MO). 2Propanol (Certified ACS) was obtained from Thermo Fisher Scientific (Waltham, MA). RIPA 10x buffer and ImmobilionTM western chemiluminescent HRP substrate were obtained from EMD Millipore (Burlington, MA). Mini-PROTEAN TGX gels and 0.45 μm nitrocellulose membranes were obtained from Bio-Rad (Hercules, CA).

Cell lines and cell culture Healthy breast epithelial cell line AG11132 was obtained from Coriell Cell Repositories (Camden, NJ). Human umbilical vein endothelial (HUVEC) cells were purchased from ATCC. Non-tumorigenic breast epithelial cell line MCF-10A and ER+ breast cancer cell line MCF-7 were purchased from ATCC. All TNBC cell lines were also from ATCC; MDA-MB-231, MDA-MB-436, MDA-MB-468, Hs578T, HCC1806, HCC1569, and DU4475. AG11132 was cultured with MEGM supplemented with insulin (5 μg/mL), hydrocortisone (0.5 μg/mL), EGF (5 ng/mL) and bovine pituitary extract (70 μg/mL). MCF10A cell line was grown with DMEM/F12 supplemented with 5 % HS, EGF (20 ng/mL), hydrocortisone (0.5 μg/mL), cholera toxin (100 ng/mL), insulin (5 μg/mL), and 1 % PS. HUVEC cells were cultured in endothelial cell growth media from R&D systems (Minneapolis, MN). MCF-7 and all the TNBC cell lines used in this study were cultured with medium and supplements specified on ATCC website. LPAR1 and G2A expression levels


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LPAR1 and G2A expression levels were quantified using a Becton Dickinson (Franklin Lakes, NJ) FACS Aria benchtop cell sorter with BD FACSDivaTM software. Briefly, HUVEC, MCF10A, and different TNBC cell lines were collected into 15 mL tubes and washed with PBS twice. Cells were then treated with 1 % BSA for 30 min to block the nonspecific binding of antibodies. Each cell line was separated into 3 different tubes and treated with PBS, FITC-conjugated rabbit polyclonal anti-EDG2 antibody and FITCconjugated rabbit polyclonal anti-G2A antibody for additional one hour in an ice bucket followed by washing with PBS twice. Cells were resuspended with fresh PBS and transferred to round-bottom polystyrene (Globe Scientific, Paramus, NJ) to perform flow cytometry analysis. Mean FITC fluorescence was compared to quantify the relative LPAR1 and G2A expression levels of each cell line. Preparation of drug encapsulating LNEs LNEs were prepared using solvent injection technique with slight modification[53]. Briefly, CLO was dissolved in 2-propanol (30 mg/mL). DAC and PAN were freshly prepared in 2-propanol : DMSO mixture (19 to 1 volume ratio) with ultrasonication in water bath at the final concentration of 0.5 mg/mL each. The dissolved epigenetic drugs and CLO were mixed and agitated for an hour as the ratio shown on Table S2. Lysophophatidylcholine (LPC), lysophophatidic acid (LPA), and DSPE-PEG (2000) carboxylic acid (DSPE-PEG-COOH) were all prepared in 1 to 1 ratio of PBS and 2-propanol at the concentration of 10 mg/mL each. Lipid components were further diluted in PBS to achieve the final mass ratio shown in Table 1, and the mixture was stirred and preheated up to 70 °C on a hot plate. Then, drug-CLO mixture in organic solvents was rapidly injected into the heated aqueous phase containing the lipids components and stirred for 7 min; the water to 2-propanol volume ratio was 10 to 1 and the total lipid mass was


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about 19 % of CLO. The heat was, then, removed from the plate and the solution kept being stirred for one more hour for the complete evaporation of 2-propanol. Finally, the fabricated LNEs were washed with PBS two times using Nanosep Centrifugal Device Omega (100 kD molecular weight cutoff) and resuspended in PBS. The first filtrate of each LNE was frozen for high performance liquid chromatography-mass spectrometry (HPLC-MS) analysis.

Blank-LNEs DAC-LNEs PAN-LNEs DAC/PAN-LNEs

Size (nm)

PDI

Zeta potential (mV)

188.0±0.7 192.4±0.5 199.3±2.2 200.5±1.0

0.118 0.170 0.136 0.103

-14.8±1.0 -12.2±0.6 -13.7±0.1 -14.7±1.8

Encapsulation Efficiency (%) 33.0±3.7 68.2±0.5 48.8±3.2(DAC) 65.1±0.7(PAN)

Table 1. Characterization of LNEs (1:1 mole ratio of LPA:LPC) Preparation of Rhodamine 123 (or DiO/DiR) encapsulating LNEs Rhodamine 123 (Rho 123) was dissolved in 2-propanol (100 μg/mL). CLO and Rho 123 were mixed at the mass ratio of 100 to 1. Rho 123 encapsulating LNEs (Rho-LNEs) were prepared using solvent injection method as described above. Three different lipid compositions were used to fabricate LNEs and the uptake efficiency was compared; 1:0, 1:1 and 0:1 mole ratio of LPA and LPC, and 5 mol % DSPE-PEG-COOH. Encapsulation of Rho 123 in different LNEs was quantified by microplate reader (SpectraMax Gemini-XPS, Molecular Devices, Sunnyvale, CA) at 510/535 wavelengths of excitation/emission after overnight dialysis in PBS. Fabrication of DiO-LNEs or DiR-LNEs were same with the above method except the addition of 0.004 % (w/v) of DiO or DiR instead of Rho 123.


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Characterization of LNEs The hydrodynamic diameters of LNEs and polydispersity index (PDI) were measured using dynamic light scattering (DLC) with BIC particle sizing software (Brookhaven Instruments Corporation, Holtsville, NY). The zeta potential of LNEs were determined in a 10mM KCl solution using BIC Pals zeta potential analyzer (Brookhaven Instrument Corporation) at 25 °C. The morphology of LNEs was observed with transmission electron microsopy (TEM, JEM-2100, JEOL Ltd, Tokyo, JAPAN). The samples were negative stained with uranyl acetate (1 % w/v) before observation. Drug loading and entrapment efficiency A standard plot for DAC (0 - 22.82 µg/mL) and PAN (0 - 0.70 µg/mL) were prepared first. The frozen samples of LNE filtrates were thawed and diluted in a mixture of organic solvents (making 50 v/v % filtrates in PBS, 25 v/v % 2-propanol, 20 v/v % acetonitrile, and 5 v/v % DMSO). DAC and PAN concentrations were determined using HPLC-MS (Dionex Ultimate-3000 U(H)PLC system, Thermo Scientific; MaXis-II ESI-TOF, Bruker, Billerica, MA). HPLC condition: Reversed-phase C18 column (3 µm, 2.1X100 mm column, Acclaim-120, Thermo Fisher Scientific) was used. The mobile phase was H2O/0.1 % formic acid (A-solvent) and acetonitrile/0.1 % formic acid (B-solvent). Gradient starts at 95 % of A–solvent and 5 % of B-solvent. Gradient at 30 min is 5 % of A-solvent and 95 % of B-solvent. Flow rate was 200 µL/min and injection volume was 4 µL. A wavelength of 245 nm was used for DAC and 214 nm for PAN using UV detector. Mass-spectrometry condition:


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MaXis-II ESI-TOF instrument was optimized to get the most sensitive conditions for the compounds of interest; capillary voltage: 4000 V, nebulizer gas: 1 L/min, curtain gas: 8 L/min, collision gas (arbitrary units): 24, heat temperature of drying gas: 180C, ion production mode: electrospray-ionization. Full-scan data was acquired by scanning from 50 to 3000 m/z. Encapsulation efficiency (EE) was obtained using the equation below. 𝐸𝐸 % =

(

) × 100 %

Weight of the drug input ― Weight of drug in the filtrate Weight of the drug input

Cell uptake study Different TNBC cell lines were seeded in 6-well plates (3×105 cells/well). After 24 h preincubation, the cells were treated with Rho-LNEs (1:0, 1:1 and 0:1 mole ratio of LPA and LPC, 71.4 ng/mL final concentration of Rho 123) in each well for 2 h. The cells were washed twice with PBS, and collected using 0.1 % trypsin/2.6 mM EDTA solution. Collected cells were further washed and resuspended in PBS. The uptake of three different Rho-LNEs in each cell line was quantified using flow cytometry. The identical experimental procedure was used for all the uptake experiment with minor changes in condition. For the antibody blocking study, MDA-MB-231 cells were pre-incubated with IgG or anti-LPAR1 for 1 h at a concentration of 10 μg/mL before treatment with DiO-LNEs. Cell uptake was analyzed at 4 °C vs 37 °C. Filipin (1.5 μM), chlorpromazine (10 μM), and dynasore (50 μM) were added to MDA-MB-231 cells 30 m prior to DiO-LNEs treatment to evaluate internalization mechanism. To visualize the uptake difference of LNEs, TNBCs were also plated on glass bottom microwell dishes (14 mm microwell, MatTek Corporation, Ashland, MA) at a concentration of 3×105 cells per well and pre-incubated for 24 h. Each cell line was treated with Rho-LNEs as described above, washed with PBS, and fixed with 4 %


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paraformaldehyde for 15 min. Cells were washed with PBS for another three times, and nuclei was stained with DAPI (5 μg/mL) for 2 min. Then, DAPI solution was removed and cells were washed, at last, for three times. The fluorescent images of cells were obtained using a confocal microscope (Zeiss LSM 710, Carl Zeiss AG, Oberkochen, Germany). Biodistribution of DiR-LNEs Primary breast cancer tumor model was constructed by injecting 5×106 of MDAMB-231 cells into the 4th mammary fat pad of athymic female nude mice (Charles River). When tumor size reached 100 -150 mm3, mice were randomized into three treatment groups; DiR-LNEs (1:0, 1:1, 0:1 mole ratio of LPA:LPC). DiR-LNEs (100 mg/kg) were intravenously injected and in vivo fluorescence was monitored at 2, 4, 6, 24, 48, and 72h post-injection using IVIS Lumina II (Caliper, Hopkinton, MA). After 72h injection,

mice were sacrificed and organs were collected and imaged ex vivo to track the organ distribution of DiR-LNEs. Drug release/stability study DAC/PAN-LNEs were added to a 10 kDa MWCO (molecule weight cut-off) dialysis tube and dialyzed against 20 mL of PBS (pH 7.4) with 0.1% (w/v) Tween 20 for 24 h. The initial PAN concentration was 1 mg/mL. 500 µL samples were collected at 0.5, 1, 2, 5, 8, and 24 h for further analysis, and supplemented with 500 µL of fresh PBS. Free DAC/PAN dissolved in PBS and DAC/PAN-LNEs in PBS were incubated at 37 °C. The initial concentrations of DAC and PAN were 0.1 mg/mL and 3.7 μg/mL regardless of formulation. 50 µL samples were taken at 0, 6, 9, and 24 h for further quantified by HPLC for DAC concentration. Cell viability assay


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Cell viability was evaluated by resazurin reduction assay. TNBCs were seeded in 96-well plates at a density of 5000 cells per well and incubated overnight. Cells were treated with PBS, DAC (5 μM), PAN (120 nM), DAC (5 μM)/PAN (120 nM), BLANK-LNEs, DAC-LNEs, PAN-LNEs, and DAC/PAN-LNEs for 48 h (equivalent concentrations of DAC and/or PAN encapsulating LNEs to free drug). The medium in each well in the plates was replaced with 90 μL of freshly warmed medium, then 10 μL of resazurin sodium salt (500 μM) was added, and incubated for 4 h. Proliferation was examined at wavelengths of 530 nm excitation and 590 nm emission using a microplate reader. Cell Growth Inhibition assay The growth inhibition assay was performed using resazurin reduction assay. MDA-MB-231 cells were plated at a density of 3×105 cells per well in 6-well plates and preincubated for 24 h. Cells were treated with free drug and LNEs as described above. After 24 h incubation, cells were harvested using 0.1 % trypsin/2.6mM EDTA solution, and seeded in a 96-well plate at a concentration of 5000 cells per well. Cells were treated with free drug and LNEs after overnight incubation. Free drug and LNEs containing medium were replenished every other day. Cell growth inhibition was examined every 24 h for 5 d using resazurin sodium salt as described above. Screening of regional methylation status of tumor suppressor genes DNA samples were collected from MDA-MB-231 cells treated with either BlankLNEs or DAC/PAN-LNEs in separate tubes using DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany). DNA digestion was performed with Epitect Methyl II DNA Restriction Kit (Qiagen) according to the manufacturer’s instruction. Briefly, 1 μg of genomic DNA was used for the following restriction enzyme digestion: (1) mock digest (Mo) (2) methylation-sensitive digest (Ms) (3) methylation-dependent digest (Md) (4) double


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digest of methylation-sensitive and methylation-dependent restriction enzymes (Msd). Each digest group was mixed with RT2 SYBR Green qPCR Mastermix (Qiagen) and added to Epitect Methyl II Complete qPCR Array. qRT-PCR was performed using Applied Biosystems (Foster City, CA) 7300 Real-Time PCR System, and acquired data was analyzed as presented in the manufacture’s handbook. The degree of methylation was calculated as follows; Unmethylated (UM) DNA fraction: 𝐹𝑈𝑀 =

𝐶𝑀𝑑 𝐶𝑀𝑜 ― 𝐶𝑀𝑠𝑑

Hypermethylated (HM) DNA fraction: 𝐹𝐻𝑀 =

𝐶𝑀𝑠 𝐶𝑀𝑜 ― 𝐶𝑀𝑠𝑑

Intermediately methylated (IM) DNA fraction: 𝐹𝐼𝑀 = 1 ― 𝐹𝐻𝑀 ― 𝐹𝑈𝑀 Expressions of CDH1 and FOXM1 in different breast cancer cells First, intrinsic levels of CDH1 and FOXM1 mRNA expression in MDA-MB-231, MCF-7, MDA-MB-436, HCC1569, and DU4475 were compared using qRT-PCR. The above cell lines were seeded into 6-well plate at a density of 3×105 cells per well and incubated for 24 h. mRNA of each cell line was collected using RNeasy Mini Kit (Qiagen) and converted to cDNA using RT2 First Strand Kit (Qiagen). cDNA from the above cell lines was mixed with optimized amounts of primer pairs of B2M, CDH1 or FOXM1 along with RT2 SYBR Green qPCR Mastermix, and Nuclease-Free water (Qiagen). B2M was used as an internal control to normalize mRNA levels between samples. Primers used are; B2M (5GATGAGTATGCCTGCCGTGT-3 and 5- GCGGCATCTTCAAACCTCC-3), CDH1 (5CTTTGACGCCGAGAGCTACA-3 and 5-AAATTCACTCTGCCCAGGACG-3), and FOXM1 (5-


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CGAGCACTTGGAATCACAGC-3 and 5-CAGGGGGAGTTCGGTTTTGA-3). The mixed samples were plated on 96-well Semi-SkirtedPCR plates (Thermo Scientific) and qRT-PCR was performed. Gene expression modulating effect of LNEs were examined similarly except that MDA-MB-231 cell line was treated with Blank-LNEs, DAC-LNEs (5 μM), PAN-LNEs (120 nM), DAC/PAN-LNEs (5 µM of DAC and 120 nM of PAN), and DOX (123 nM, IC50) for 20 h before mRNA was collected from each group. E-cadherin expressions in different breast cancer cells E-cadherin expression levels were quantified and compared using flow cytometry. First, intrinsic E-cadherin expression levels of MDA-MB-231, MCF-7, MDA-MB436, HCC1569, and DU4475 were compared. Each cell line was seeded on 6-well plates at a density of 3×105 cells per well and incubated for 24 h. Each cell line was washed with PBS, trypsinized, and collected at a density of 3×105 cells per tube. Each tube was further washed with PBS and incubated with 1 % BSA for 30 min to block the nonspecific binding followed by 1 hour incubation with isotype control IgG or mouse anti-human CD324. Samples were washed with PBS, resuspended in PBS, and intrinsic E-cadherein expression levels of the above cell line were evaluated using flow cytometry. Mean APC fluorescence of each cell line was compared for the relative E-cadherin expression levels. Similarly, E-cadherin expression changes of MDA-MB-231 were examined using flow cytometry after 24 h treatment of Blank-LNEs and DAC/PAN-LNEs. Western blot analysis Lysates of MDA-MB-231 treated with different LNEs were collected using lysis buffer (RIPA buffer, protease inhibitor, phosphatase inhibitor, ASONOV, and PMSF). Proteins were separated on Mini-PROTEAN TGX precast gels using electrophoresis, and


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transferred to nitrocellulose membrane. Membranes were incubated with HPRT1 (1:2000) or FOXM1 (1:1000) overnight on an orbital shaker followed by 1 hr incubation with Amersham ECL Rabbit IgG antibody (NA934V, 1:10,000) or m-IgGĸ BP(sc-516102, 1:2000). Bands were visualized with ImmobilionTM western chemiluminescent HRP substrate and quantified using Image Lab software(Bio-Rad). Statistical analysis Obtained data was compared using One-way ANOVA (α=0.05) with post-hoc Tukey test and Student’s t-test. Error bars in the figures are the standard deviation for a minimum of n=3, referring to replicates within the same experiment. Each formulation was synthesized three times and characterized independently.


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Results LPAR1 and G2A expression levels in TNBCs LPAR1 and G2A expression were quantified in TNBC cells (Figure 1A). LPAR1 expression was gauged as high (between 4.4 and 5.5–fold higher) in HCC1569, DU4475, and MBA-MD-468 TNBC cell lines; moderate (between 2.7 and 3.2-fold) in HS578T, MDAMB-436, and HCC1806 TNBC cell lines; and low in MDA-MB-231 (2.0-fold) relative to nontumorigenic MCF10A and human umbilical vein endothelial cells (HUVECs). LPC, used as a control ligand, is the precursor of LPA; its target receptor, G2A, was also measured. TNBCs showed only 1.2 to 2.1-fold greater expression of G2A in comparison to HUVEC. Pronounced LPAR1 upregulation in TNBCs compared to MCF10A and HUVEC cells suggested that LPAR1 may be utilized as a molecular target for TNBCs. LPA and LPC activity LPA may be useful to target the LPAR1 receptor but it may also increase cell metabolism. To understand the proliferative effect of LPA and LPC on TNBC cell lines, we measured cell growth over a period of 48 h as a function of concentration. Figure 1B and 1C demonstrated that LPA and LPC did not significantly promote the proliferation of TNBC cell lines compared to the sham control group.


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Figure 1. LPAR1 and G2A expression and activity in TNBC cell lines. (A) Different cell lines were treated with FITC-conjugated anti-EDG2 (LPAR1 antibody) and anti-G2A antibody (70 µg/mL). The expression levels of LPAR1 and G2A were measured using flow cytometry. Represented statistical significance applies only to LPAR1. Proliferation of TNBCs were measured after 48 h of incubation with either LPA (B) or LPC (C) using resazurin reduction assay. Error is 1 standard deviation of the mean. Significance was determined by Oneway Anova and Tukey post hoc test were performed (n=3, * p < 0.01).

Preparation and characterization of dye encapsulating LNEs To optimize the uptake efficiency of LNEs, three different LNEs were prepared from mixtures of LPA and LPC, as shown in Table S1, using a modified solvent injection method as described in Materials and Methods. To evaluate uptake efficiency, Rhodamine 123 (Rho123) encapsulating LNEs (Rho-LNEs) were prepared. Rho-LNEs were comprised of Rho123 solubilized in a CLO core, 5 mol % DSPE-PEG-COOH, and LPA and LPC at either 1:0, 1:1, or 0:1 molar ratio. The diameter of Rho-LNEs were 183.5 ± 1.0, 197.1 ± 1.8, 187.3 ± 1.2 for 1:0, 1:1, and 0:1 ratios of LPA:LPC respectively (Table S1).

In vitro LNE uptake Uptake by TNBC cell lines was measured after 2 h of incubation with Rho-LNEs with three different LPA:LPC ratios (1:0,1:1, and 0:1). Flow cytometry analysis showed that TNBC cell lines with high LPAR1 expression (Figure 1A) had higher uptake of RhoLNEs with 1:1 or 1:0 LPA:LPC (Figure 2A). Confocal images confirmed dependence of RhoLNEs uptake as a function of LPA:LPC ratio on LPAR1 expression (Figure S1). Low fluorescence was observed in the cytoplasm of MDA-MB-231 cells, having low LPAR1


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expression while high fluorescence was detected in the cytoplasm of MDA-MB-468, having high LPAR1 expression. TNBC cell lines with intermediate levels of LPAR1 expression showed a similar trend of Rho-LNE uptake. Rho-LNEs with a 1:1 ratio of LPA:LPC had 5.4 to 19.2 times higher uptake compared to 0:1 LPA:LPC Rho-LNEs. LPA:LPC Rho-LNEs with a 1:0 ratio exhibited 4.5 to 13.4 higher uptake relative to 0:1 LPA:LPC RhoLNEs. Interestingly, 1:1 LPA:LPC LNEs showed equal or higher uptake relative to 1:0 LPA:LPC LNEs. Due to the high uptake, LNEs with 1:1 LPA:LPC was chosen as a delivery vehicle for DAC and PAN. MDA-MB-231 cells were incubated with DiO encapsulating LNEs (1:1, LPA:LPC) at 4 or 37 °C to confirm the receptor mediated uptake of LNEs (Figure S2A, S2B). Confocal microscopy and flow cytometry analysis demonstrated a significantly higher uptake of DiO-LNEs at 37 °C relative to 4 °C. To demonstrate the specificity of LPAR1 targeting LNEs, MDA-MB-231 cells were blocked with IgG or anti-LPAR1 before treatment with DiO-LNEs dissolved in ethanol (Solution) or DiO-LNEs (Figure 2B, S3). Flow cytometry and confocal microscopy data showed that the Solution group had low uptake of DiO regardless of IgG or antibody treatment. Uptake of DiO-LNEs was significantly impaired when the LPAR1 antibody (antiLPAR1) was added relative to the IgG treated group. Thus, LNEs are dependent on LPAR1 for binding and uptake. Caveolae, clathrin, and both caveolae/clathrin endocytosis were blocked with filipin, chlorpromazine, and dynasore, respectively (Figure S4). Blockage of caveolae did not affect uptake of DiO-LNEs. When clathrin was inhibited, uptake of DiO-LNEs was lowered by 12 %. Uptake of DiO-LNEs decreased by 70 % when both caveolae/clathrin


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were blocked. This confirms that both caveolae and clathrin are involved in internalization of LNEs.

In vivo biodistribution by TNBC cells To examine the biodistribution of LNEs, in vivo imaging of mice and ex vivo imaging of organs were conducted. In vivo imaging showed higher accumulation of DiRLNEs (1:1, LPA:LPC) in TNBC tumors than that of DiR-LNEs (1:0 and 0:1, LPA:LPC) (Figure 2C). The ex vivo DiR signal was used to quantify the distribution of LNEs in different organs relative to surface area. Liver accumulation of DiR-LNEs (1:0, LPA:LPC) was the highest. However, it was reduced upon introduction of LPC in the LNE formulation (Figure 2D). DiR signal in tumors confirmed higher accumulation of DiR-LNEs (1:1, LPA:LPC) compared to DiR-LNEs (1:0, LPA:LPC) accumulation (Figure 2E, 2F). Retention of DiR-LNEs in mice (1:0, 1:1, and 0:1 ratio of LPA and LPC) was compared by measuring the whole body radiance of DiR-LNEs (Figure S5). DiR-LNEs (1:1, LPA:LPC) had higher residence time compared to DiR-LNEs (1:0 and 0:1, LPA:LPC) after 24 h injection.


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Figure 2. Uptake and biodistribution of LNEs (A)TNBC cell lines expressing different levels of LPAR1 were incubated with Rho-LNEs (1:0, 1:1, and 0:1 of LPA and LPC each) for 2 h.


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Fluorescence intensity was measured by flow cytometry. (B) MDA-MB-231 cells were preincubated with IgG or anti-LPAR1 at a concentration of 10 μg/ml for 1 h before incubation with DiO-LNEs (1:1, LPA:LPC). Cell uptake of LNEs was measured using flow cytometry. (C) Whole body DiR-LNEs radiance were monitored using IVIS for 72 hrs after IV injection. (D) Organ distribution of DiR-LNEs (1:0, 1:1, and 0:1 of LPA and LPC) was quantified ex vivo using IVIS 72 hrs after IV injection. (E,F) Radiance from tumors treated with DiR-LNEs. Significance was determined by One-way Anova followed by Tukey post hoc test was performed or Fisher’s Least Significant Difference post hoc test (n=3, * p < 0.05, ** p < 0.01, *** p < 0.001).


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Figure 3. Cytotoxicity of free drug and LNEs in TNBC cell lines. Non-neoplastic AG11132 cells (A) and TNBC MDA-MB-231 cells (B) were treated with PAN or PAN+DAC for 48 h. DAC concentration was fixed at 5 µM. Cell viability was measured using resazurin reduction assay (C) Cytotoxicity of free drug and LNEs (1:1 mole ratio of LPA:LPC) were compared in HUVEC, MCF10A and MDA-MB-231 after 48 h of treatment using the resazurin reduction assay. Cell viability of HCC1806 (D) and MDA-MB-468 (E) after 48 h treatment of free drugs and LNEs (1:1, LPA:LPC). (F) Proliferation of MDA-MB-231 were observed under the same conditions for 120 h d using resazurin reduction assay. Medium and drugs were replenished every two days. Error is 1 standard deviation of the mean. Significance was determined by One-way Anova and Tukey post hoc test were performed (n=3, * p < 0.05, ** p < 0.01, *** p < 0.001).


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Combinatorial effect of DAC and PAN on MDA-MB-231 growth Normal epithelial (AG11132) and TNBC (MDA-MB-231) cells were used to evaluate the dose response of PAN in the presence of 5 μM DAC. DAC alone did not induce cytotoxicity in MDA-MB-231 (up to 32 μM) after 48 h (Figure S6). The concentration of DAC (5 μM) was chosen based on prior studies where DAC treatment showed a therapeutic benefit in leukemia and a maximal steady state plasma level in clinical trials[54, 55]. The cell viability of AG11132 cells dropped by 40 % when treated with 100 nM PAN alone and did not decrease at higher concentrations (Figure 3A-B). In contrast, co-treatment of PAN and DAC reduced MDA-MB-231 cell viability by 60 %. The combination of 5 μM DAC and 100nM PAN were chosen for subsequent studies due to the synergistic toxicity on TNBCs, while minimizing the effect on AG11132 cells.

Preparation and characterization of DAC/PAN encapsulating LNEs DAC and/or PAN were formulated into LNEs (1:1 LPA:LPC) that target the LPAR1 receptor on TNBC cells (Figure S7, Table S2). TEM images confirmed a homogeneous size distribution of LNEs with a spherical shape (Figure S8). The characterization of LNEs is summarized in Table 1. The diameter of DAC/PAN-LNEs remained stable for 48 h of incubation in cell culture medium supplemented with 50 % mouse plasma (Figure S9). The stability of DAC encapsulated in DAC/PAN LNEs was measured. The active form of DAC was evaluated in a solution of free DAC/PAN or DAC/PAN-LNEs as a function of time at 37 °C (Figure S10A, S10B). Eighty percent of free DAC was degraded in water at 24 h post-incubation compared to 40 % of DAC encapsulated in LNEs. PAN release was also measured in 0.1% Tween 20 supplemented PBS (pH 7.4) at 37 °C (Figure S10C). Only


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15 % of PAN was released over 24 h, indicating the high retention of PAN within DAC/PAN-LNEs. LNEs retain PAN during circulation and maintain DAC activity. Co-delivery of DAC and PAN by LNEs The cytotoxicity of free drug (DAC, PAN, or DAC/PAN) and drug-encapsulating LNEs (Figure 3C) were examined in HUVEC (endothelial), MCF10A (nonneoplastic) and MDA-MB-231 (TNBC) cells with the concentrations determined from the previous study (5 μM DAC and 120 nM PAN, Figure S6, 3B). Due to the encapsulation efficiency, the concentration of DAC and PAN were adjusted to maintain 5 μM DAC. DAC alone was not cytotoxic. Free PAN and DAC/PAN showed a mild effect on the cell viability of MCF10A cells. In contrast, HUVEC cells were almost eradicated by free PAN and DAC/PAN. Free PAN and DAC/PAN decreased the cell viability of MDA-MB-231 by 58 % and 61 %, respectively. LNEs showed a similar trend. Blank-LNEs and DAC-LNEs were not effective in killing HUVEC, MCF10A and MDA-MB-231 cells. PAN-LNEs did not exhibit toxicity in HUVEC cells, and were less potent in MCF10A and MDA-MB-231 cells. DAC/PAN-LNEs reduced cell viability of HUVEC, MCF10A, and MDA-MB-231 cells by 24 %, 23 %, and 55%, demonstrating the superior therapeutic efficacy in TNBCs. Toxicity of DAC/PAN and DAC/PAN-LNEs was not statistically significant in MDA-MB-231 cells. A similar or greater therapeutic effect was observed in other TNBC cell lines (Figure 3D, 3E, S12). Cell growth of MDA-MB-231 cells was measured after treatment with free drug or LNEs for 5 d (Figure 3F). Blank-LNEs had no toxicity. Free DAC decreased cell viability by 12 %, while DAC-LNEs repressed cell growth by 30 % compared to the control at day 5. PAN and PAN-LNEs significantly decreased cell viability by 88 % and 84 %, respectively. The combination of DAC/PAN and DAC/PAN-LNE treatment induced maximum cell growth inhibition by up to 94 % and 92 %, respectively.


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Figure 4. Effect of DAC/PAN-LNEs (1:1, LPA:LPC) on methylation of CDH1 promoter region and gene/protein expression. (A) MDA-MB-231 cells were treated with either Blank-LNEs or


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DAC/PAN-LNEs for 20 h and methylation status of promoter regions were screened using Epitect Methyl II Complete qPCR Array. Changes of promoter methylation status greater than 10 % presented. (B) Complete demethylation of intermediately methylated (IM) CDH1 promoter region was identified. (C) Intrinsic CDH1 expression in MCF-7 and MDA-MB-231 were measured using qRT-PCR. (D) MDA-MB-231 were treated with LNEs for 20 h and changes of CDH1 and E-cadherin expression level were measured using qRT-PCR and flow cytometry each. (E) Intrinsic FOXM1 expression in MCF-7 and MDA-MB-231 were measured using qRT-PCR (F) FOXM1 mRNA and protein expression were compared after 20 h treatment of LNEs in MDA-MB-231 using qRT-PCR and Western Blot each. Concentration of DAC and PAN were 5 µM and 120 nM. Error is 1 standard deviation of the mean. Significance was determined by Student’s T-test and One-way Anova with Tukey post hoc test were performed (n=3, * p < 0.05, ** p < 0.01, *** p < 0.001,).

***


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Verification of DAC/PAN function To verify that DAC/PAN induced a change in cancer epigenetics, we used a PCR array to measure the methylation status change in tumor suppressor gene promoters by comparing MDA-MB-231 treated with Blank-LNEs and DAC/PAN-LNEs (Table S3). The promoter regions were either hypomethylated or hypermethylated after 20 h of treatment (Figure 4A, Figure S13). Among these genes, the methylation status of EpCAM, RARB, RRAD, and CDH1 promoter regions changed by more than 40 %. Absence of a key epithelial marker CDH1 in mesenchymal TNBCs is observed and associated with poorer prognosis. Thus, CDH1 was chosen to be further analyzed. We measured CDH1 mRNA and the protein E-cadherin it encodes before and after LNE treatment. The intrinsic levels of CDH1 and E-cadherin in MDA-MB-231 cells were significantly lower than those of MCF-7 cells (Figure 4C, S14). However, complete demethylation of intermediate methylated (IM) CDH1 promoter regions was observed in MDA-MB-231 cells after DAC/PAN-LNEs treatment (Figure 4B), which increased CDH1 by 3.8-fold and E-cadherin expression by 2-fold (Figure 4D). In contrast, DAC-LNE treatment had a negligible effect on CDH1 expression in MDA-MB-231 cells, suggesting that hypomethylation of the CDH1 promoter region may be mediated by PAN. We confirmed that DAC/PAN-LNE treatment reactivated CDH1 and E-cadherin expression by hypomethylating the promoter region of CDH1 in MDA-MB-231. Suppression of FOXM1 by DAC/PAN-LNEs We also measured the expression of oncogene forkhead box M1 (FOXM1), which plays an important role in the epithelial to mesenchymal transition (EMT) and chemoresistance.[56] Intrinsic FOXM1 mRNA expression of MDA-MB-231 was 40 % higher than MCF-7 (Figure 4E); this is the opposite trend of what was measured for CDH1.


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MDA-MB-231 cells were treated with LNEs encapsulating DAC, PAN, or DAC/PAN. DAC/PAN-LNEs synergistically reduced FOXM1 mRNA and FOXM1 protein expressions up to 80 % (Figure 4F, S15).

Biomarkers for DAC/PAN-LNEs treatment To determine if DAC/PAN LNE toxicity was dependent on CDH1/FOXM1 expression, we examined CDH1/E-cadherin and FOXM1 expression in four breast cancer cell lines. Mesenchymal TNBC cell lines (MDA-MB-231 and MDA-MB-436) were toxic to DAC/PAN LNEs relative to epithelial-like TNBC cells (DU4475) and HER2+ breast cancer cells (HCC1569) (Figure 5A). HCC1569 and DU4475 cells expressed significantly higher CDH1/Ecadherin and lower FOXM1 than mesenchymal TNBCs (Figure 5B-D). We investigated whether CDH1 and FOXM1 expression correlated with relapse free survival (RFS) of TNBC patients. TNBC samples from 149 patients were classified by CDH1 and FOXM1 expression. We confirmed that high FOXM1 and low CDH1 correlated with decreased RFS in TNBCs (Figure 5E-F). Overall, our results demonstrate that DAC/PAN-LNEs preferentially induced cytotoxicity in breast cancers with a CDH1(low)/FOXM1(high) expression profile, which is associated with lower RFS in TNBCs.


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Figure 5. CDH1/FOXM1 expression as a biomarker for DAC/PAN-LNEs (1:1, LPA:LPC) toxicity and its clinical association. (A) Cytotoxicity of LNEs was measured in TNBC cell


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lines and a HER2+ cell line (HCC1569) after 48 h of treatment. Concentration of DAC and PAN were 5 µM and 120 nM. Intrinsic CDH1 (B) and FOXM1 (D) expression in TNBCs were measured using qRT-PCR, and E-cadherin expression was measured using flow cytometry (C). Error is 1 standard deviation of the mean. Significance was determined by One-way Anova and Tukey post hoc test was performed (n=3, * p < 0.05, ** p < 0.01, *** p < 0.01). Relapse free survival Kaplan-Meier curves of 149 invasive breast cancers were classified by CDH1 (E) and FOXM1 (F) expression through PROGgeneV2. Gene expression data of CDH1 and FOXM1 was extracted from Gene Expression Omnibus (GSE16987).


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Discussion Recent advancements in cancer epigenetics revealed that abnormal chromatin structure plays a significant role in cancer progression by modulating gene expression[57]. Epigenetic therapy is shown to have the capability to reverse abnormal epigenetics in cancer cells with reduced toxicity to normal cells, providing an alternative to conventional chemotherapeutics[58-60]. In this study, we aimed to develop a targeted drug delivery system that could co-deliver a DNMTi and a HDACi to reverse cancer epigenetics and to induce cytotoxicity within TNBCs. LNEs (with different ratios of LPA:LPC) were fabricated to target LPAR1 on TNBCs. We identified elevated expression of LPAR1 in TNBC cell lines relative to normal cells (Figure 1A). An optimal 1:1 ratio of LPA:LPC exhibited greater binding in most TNBC cell lines relative to the 1:0 and 0:1 ratio in vitro (Figure 2A, Figure S1). DSPE- PEG was incorporated because it is known to improve the pharmacokinetics of nanoparticles by hindering protein adsorption and opsonization[61, 62]. Maximum tumor cell uptake of LNEs (1:1, LPA:LPC) was also validated by ex vivo imaging of tumors collected from DiRLNEs treated mice (Figure 2E, 2F). In order to understand the clearance mechanism and off-target effects, the biodistribution of LPAR1 targeted LNEs was assessed. Due to the stability of LNEs in 50% plasma and low release of drug, we expect that DAC and PAN would be distributed in the vicinity of LNEs or intracellularly. LNEs exhibited relatively higher accumulation in liver and intestines relative to other organs. Introduction of LPC in LNE formulation significantly reduced liver accumulation and increased tumor accumulation of LNEs. There are two reasons that may induce higher accumulation of DiR-LNEs (1:0, LPA:LPC) in liver in comparison to DiR-LNEs (1:1 and 0:1, LPA:LPC). First, liver is the primary


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clearance organ. Hepatocytes, Kupffer cells, liver sinusoidal endothelial cells (LSECs), motile macrophage cells, hepatic stellar cells, biliary epithelial cells, and resident immune cells all interact together to capture and clear foreign molecules[63, 64]. Interestingly, Kupffer cells and LSECs abundantly express anion-scavenger receptors on their membranes[65]. The highly negatively charged DiR-LNEs (1:0, LPA:LPC) may be preferentially filtered in the liver by Kupffer cells and LSECs. Second, LPA receptors are ubiquitously expressed in immune cells and LPA stimulates immune cells[66]. Stimulation of immune cells in the liver by DiR-LNEs (1:0, LPA:LPC) may activate the immune system, thereby leading to more clearance from liver. In the intestines, LPAR1 is important in maintaining epithelial barrier integrity[67]. It is not surprising that LNEs would accumulate in the intestines due to their LPAR1 expression. Cells in the intestines have a high turnover rate and often respond to chemotherapeutics [68, 69]. DAC/PAN LNEs, however, have lower toxicity in normal cells than TNBC cells. Blood circulation of nanoparticles is dependent on several factors such as size, surface charge, and surface chemistry [70]. Considering the significantly lower liver accumulation of DiR-LNEs (1:1, LPA:LPC) compared to DiR-LNEs (1:0, LPA:LPC) and relatively minimal accumulation of LNEs in other organs, there may be a higher amount of DiR-LNEs (1:1, LPA:LPC) in the blood stream relative to DiR-LNEs (1:0, LPA:LPC) (Figure 2D, S6). Increased retention of DiR-LNEs may be due to the slightly negative surface charge from the carboxyl group in LPA, the presence of zwitterionic phospholipids (LPC), and PEG shielding[71]. During circulation or within the tumor environment, PEG-DSPE disassociate from LNEs and may associate within other lipids or hydrophobic proteinss. Taken together, LNEs, less than 200 nm in diameter, negatively charged, and shielded by PEG were used to actively target a lipid binding receptor.


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The dependence of the LPA:LPC ratio on uptake efficiency may be attributed to two reasons. First, the increase of LPA in LNEs decreases the negative surface potential and improves stability (Table S1). Second, LNE uptake is mediated by LPAR1 since the uptake of LNEs in TNBCs is LPAR1 expression level dependent. Receptor-mediated endocytosis requires ATP[72, 73]. Impaired uptake of DiOLNEs at 4 °C in comparison to 37 °C suggests that uptake of LNE is mediated by LPAR1 (Figure S2). LPAR1 mediated uptake of LNEs was confirmed by reduced uptake of DiOLNEs when LPAR1 was blocked with anti-LPAR1 (Figure 2B, S3). LPAR1 is expressed in adult tissues and immune cells[74]. LPAR1 expression is commensurate with stage of breast and ovarian cancers[75, 76]. Increased blood levels of LPA in aggressive cancers suggest increased LPAR receptor expression and activity[47]. Thus, in spite of relatively low levels of LPAR1 in MDA-MB-231 cells in vitro, LPAR1 may have higher expression and activity in tumors in vivo, with dependence on tumor progression. Thus, our platform may have enhanced targeting efficiency for late stage tumors. This hypothesis needs further evaluation. A previous study showed that LPAR1 is co-localized with caveolae/clathrin on the plasma membrane[77]. The endocytic pathway inhibition experiment confirmed that LPAR1-mediated endocytosis is mediated by both caveolae and clathrin (Figure S4). The LPA-LPAR1 axis was previously reported to accelerate cancer cell proliferation in a serum starved condition[78]. In our study, LPA and LPC did not promote TNBC proliferation in 10 % serum (Figure 1B-C). Blank-LNEs did not promote the proliferation of MCF-10A and MDA-MB-231 (Figure S11). Thus, LPA and LPC did not contribute to cell growth or toxicity.


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DAC and PAN, in combination, regardless of the formulation, were cytotoxic to a variety of TNBC cell lines, while non-neoplastic breast cells showed less toxicity (Figure 3, 5A, S12). PAN-LNEs and DAC/PAN-LNEs were more potent or as effective as free drug in TNBCs. Interestingly, PAN toxicity was abrogated in HUVEC cells when encapsulated in LNEs likely due to the slow release of PAN and low uptake (Figure S10C and 1A), which may inhibit cells from reaching an effective drug concentration. PAN was previously shown to kill TNBC cell lines[20]. Our results also exhibited potent cytotoxicity of PAN in TNBC cell lines (Figure 3, Figure S12). DAC undergoes hydrolytic decomposition within hours at physiological pH and temperature[79-81]. Past studies demonstrated the enhanced regulatory effect of DAC on cancer cells when it is delivered through a drug delivery carrier, indirectly showing the improved stability of DAC by limiting hydrolytic and/or enzymatic degradation[82, 83]. We showed that LNEs significantly delayed hydrolysis of DAC in comparison to free DAC (Figure S10A, S10B). Interestingly, DAC-LNEs were more potent compared to free DAC in cell growth inhibition, indicating that improved stability of DAC may have a therapeutic benefit in TNBCs (Figure 3F). The effect of DAC/PAN-LNEs on gene and protein expression showed a performance benefit that may affect cancer progression. CDH1 in primary breast cancers is often reduced even though no CDH1 mutation is found[84]. This may be due to hypermethylation of the promoter region of CDH1 along with other epigenetic abnormalities, which contributes to the reduced expression of CDH1 and E-cadherin[85]. Loss of E-cadherin is associated with tumorigenesis, EMT, and poor prognosis in a variety of cancers[86-90]. Reactivation of CDH1/E-cadherin in TNBCs repressed cell migration in vitro[91-93]. In our study, DAC/PAN-LNE treatment completely demethylated the IM


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CDH1 promoter regions, leading to increased expression of CDH1 and E-cadherin in MDAMB-231 cells (Figure 4B, 4D). DAC/PAN-LNEs inhibited the migration of MDA-MB-231 by 60 % in comparison to sham control group (Figure S16). FOXM1 is considered a regulator of tumor metastasis, angiogenesis, proliferation, migration and chemoresistance[56]. Previous studies demonstrated that FOXM1 expression is downregulated by long term treatment with DAC at a low dosage (20 mg/m2, five times a week for five weeks) in leukemia patients, and by PAN treatment in gastric cancer cell lines Thus, we hypothesized that FOXM1 expression in TNBCs may also be modulated by DAC/PAN-LNEs. DAC/PAN treatment inhibited the expression of FOXM1 mRNA and protein in MDA-MB-231 cells (Figure 4F, S15). Interestingly, the inverse relationship of a key epithelial marker CDH1/E-cadherin relative to FOXM1 expression was also observed before and after DAC/PAN-LNEs treatment (Figure 5B-D). The overexpression of FOXM1 and downregulation of E-Cadherin are strongly correlated with poor prognosis of breast cancers[75] [94]. Silencing of FOXM1 in MDA-MB-231 cells resulted in the downregulation of mesenchymal markers and upregulation of epithelial markers while forced expression of FOXM1 in MCF-7 cells showed an opposite trend in EMT marker expressions[95]. This shows that DAC/PAN LNEs can increase tumor suppressor genes and decrease oncogenes.

Conclusion We synthesized LPAR1-targeted LNEs encapsulating DAC and PAN to selectively induce cytotoxicity to TNBC cells. For the first time, we showed that LPAR1 overexpressing TNBCs can be targeted using LPA/LPC LNEs in vitro and in vivo. Extended blood circulation of LPAR1-targeted LNEs (1:1, LPA:LPC) may enable targeted drug


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delivery to TNBC tumors. DAC/PAN delivery through LNEs restored CDH1/E-cadherin expression and suppressed FOXM1 expression in MDA-MB-231, leading to a decrease in cell viability. We also showed that DAC/PAN-LNEs selectively killed CDH1(low)/FOXM1(high) TNBCs. Overall, our study showed that LPAR1 targeted, DAC/PAN LNEs may be a therapeutic alternative to chemotherapeutics for mesenchymal TNBCs.

Acknowledgements I would like to thank CUNY Advanced Science Research Center for mass spectral data, RCMI and CCNY Electron Microscopy Center for allowing me to use flow cytometry, confocal microscope, and TEM. This work was supported by the National Institutes of Health [DP2-CA174495-01].

Conflicts of interest: None

Supporting Information Supporting Information is available. Characterization of LNEs, a list of genes evaluated by microarray, LNE uptake images of confocal microscopy, whole body radiance of LNE, cytotoxicity of LNEs, cell migration, Ecadherin/FOXM1 expression


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Reference 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

Weigelt, B., et al., Refinement of breast cancer classification by molecular characterization of histological special types. J Pathol, 2008. 216(2): p. 141-50. Weigelt, B. and J.S. Reis-Filho, Histological and molecular types of breast cancer: is there a unifying taxonomy? Nat Rev Clin Oncol, 2009. 6(12): p. 718-30. Anders, C.K. and L.A. Carey, Biology, metastatic patterns, and treatment of patients with triplenegative breast cancer. Clin Breast Cancer, 2009. 9 Suppl 2: p. S73-81. Haffty, B.G., et al., Locoregional relapse and distant metastasis in conservatively managed triple negative early-stage breast cancer. J Clin Oncol, 2006. 24(36): p. 5652-7. Dent, R., et al., Triple-negative breast cancer: clinical features and patterns of recurrence. Clin Cancer Res, 2007. 13(15 Pt 1): p. 4429-34. Saibil, S., et al., Incidence of taxane-induced pain and distress in patients receiving chemotherapy for early-stage breast cancer: a retrospective, outcomes-based survey. Current Oncology, 2010. 17(4): p. 42-47. Azim, H.A., et al., Long-term toxic effects of adjuvant chemotherapy in breast cancer. Annals of Oncology, 2011. Swain , S.M., et al., Longer Therapy, Iatrogenic Amenorrhea, and Survival in Early Breast Cancer. New England Journal of Medicine, 2010. 362(22): p. 2053-2065. Feinberg, A.P., R. Ohlsson, and S. Henikoff, The epigenetic progenitor origin of human cancer. Nat Rev Genet, 2006. 7(1): p. 21-33. Jenuwein, T. and C.D. Allis, Translating the Histone Code. Science, 2001. 293(5532): p. 1074-1080. Feinberg, A.P. and B. Tycko, The history of cancer epigenetics. Nature Rev. Cancer, 2004. 4: p. 143153. Sharma, S., T.K. Kelly, and P.A. Jones, Epigenetics in cancer. Carcinogenesis, 2010. 31(1): p. 27-36. Jones, P.A. and S.B. Baylin, The Epigenomics of Cancer. Cell, 2007. 128(4): p. 683-692. Reid, G.K., J.M. Besterman, and A.R. MacLeod, Selective inhibition of DNA methyltransferase enzymes as a novel strategy for cancer treatment. Curr Opin Mol Ther, 2002. 4(2): p. 130-7. Creusot, F., G. Acs, and J.K. Christman, Inhibition of DNA methyltransferase and induction of Friend erythroleukemia cell differentiation by 5-azacytidine and 5-aza-2'-deoxycytidine. J Biol Chem, 1982. 257(4): p. 2041-8. Fraga, M.F., et al., Loss of acetylation at Lys16 and trimethylation at Lys20 of histone H4 is a common hallmark of human cancer. Nat Genet, 2005. 37(4): p. 391-400. Ceccacci, E. and S. Minucci, Inhibition of histone deacetylases in cancer therapy: lessons from leukaemia. British Journal of Cancer, 2016. 114(6): p. 605-611. Carrier, F., Chromatin Modulation by Histone Deacetylase Inhibitors: Impact on Cellular Sensitivity to Ionizing Radiation. Mol Cell Pharmacol, 2013. 5(1): p. 51-59. Bhattacharjee, D., S. Shenoy, and K.L. Bairy, DNA Methylation and Chromatin Remodeling: The Blueprint of Cancer Epigenetics. Scientifica, 2016. 2016: p. 11. Tate, C.R., et al., Targeting triple-negative breast cancer cells with the histone deacetylase inhibitor panobinostat. Breast Cancer Res, 2012. 14(3): p. R79. Xia, C., et al., Treatment of resistant metastatic melanoma using sequential epigenetic therapy (decitabine and panobinostat) combined with chemotherapy (temozolomide). Cancer Chemotherapy and Pharmacology, 2014. 74(4): p. 691-697. Fiskus, W., et al., Panobinostat treatment depletes EZH2 and DNMT1 levels and enhances decitabine mediated de-repression of JunB and loss of survival of human acute leukemia cells. Cancer Biology & Therapy, 2009. 8(10): p. 939-950. Kalac, M., et al., HDAC inhibitors and decitabine are highly synergistic and associated with unique gene-expression and epigenetic profiles in models of DLBCL. Blood, 2011. 118(20): p. 5506-5516. Luszczek, W., et al., Combinations of DNA Methyltransferase and Histone Deacetylase Inhibitors Induce DNA Damage in Small Cell Lung Cancer Cells: Correlation of Resistance with IFN-Stimulated Gene Expression. Molecular Cancer Therapeutics, 2010. 9(8): p. 2309-2321.


41


25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46.

Page 42 of 45

Rogstad, D.K., et al., The Chemical Decomposition of 5-aza-2′-deoxycytidine (Decitabine): Kinetic Analyses and Identification of Products by NMR, HPLC, and Mass Spectrometry. Chemical research in toxicology, 2009. 22(6): p. 1194-1204. Jabbour, E., et al., Evolution of decitabine development: accomplishments, ongoing investigations, and future strategies. Cancer, 2008. 112(11): p. 2341-51. Hennika, T., et al., Pre-Clinical Study of Panobinostat in Xenograft and Genetically Engineered Murine Diffuse Intrinsic Pontine Glioma Models. PLOS ONE, 2017. 12(1): p. e0169485. Younes, A., et al., Panobinostat in Patients With Relapsed/Refractory Hodgkin's Lymphoma After Autologous Stem-Cell Transplantation: Results of a Phase II Study. Journal of Clinical Oncology, 2012. 30(18): p. 2197-2203. Shah, P., D. Bhalodia, and P. Shelat, Nanoemulsion: A Pharmaceutical Review. Systematic Reviews in Pharmacy, 2010. 1(1): p. 24-32. Sonneville-Aubrun, O., J.T. Simonnet, and F. L'Alloret, Nanoemulsions: a new vehicle for skincare products. Advances in Colloid and Interface Science, 2004. 108–109: p. 145-149. Sanders, T.A., M. Vickers, and A.P. Haines, Effect on blood lipids and haemostasis of a supplement of cod-liver oil, rich in eicosapentaenoic and docosahexaenoic acids, in healthy young men. Clin Sci (Lond), 1981. 61(3): p. 317-24. Simopoulos, A.P., The importance of the ratio of omega-6/omega-3 essential fatty acids. Biomedicine & Pharmacotherapy, 2002. 56(8): p. 365-379. Guil-Guerrero, J.L. and E.-H. Belarbi, Purification process for cod liver oil polyunsaturated fatty acids. Journal of the American Oil Chemists' Society, 2001. 78(5): p. 477-484. Skeie, G., et al., Cod liver oil, other dietary supplements and survival among cancer patients with solid tumours. International Journal of Cancer, 2009. 125(5): p. 1155-1160. Dyck, M.C., D.W.L. Ma, and K.A. Meckling, The anticancer effects of Vitamin D and omega-3 PUFAs in combination via cod-liver oil: One plus one may equal more than two. Medical Hypotheses, 2011. 77(3): p. 326-332. Garland, C.F., et al., The Role of Vitamin D in Cancer Prevention. American Journal of Public Health, 2006. 96(2): p. 252-261. Chime, S.A., F.C. Kenechukwu, and A.A. Attama, Nanoemulsions — Advances in Formulation, Characterization and Applications in Drug Delivery, in Application of Nanotechnology in Drug Delivery, A.D. Sezer, Editor. 2014, InTech: Rijeka. p. Ch. 03. Liu, D. and D.T. Auguste, Cancer targeted therapeutics: From molecules to drug delivery vehicles. Journal of Controlled Release, 2015. 219: p. 632-643. Guo, P., et al., ICAM-1 as a molecular target for triple negative breast cancer. Proceedings of the National Academy of Sciences, 2014. 111(41): p. 14710-14715. Veneti, E., R.S. Tu, and D.T. Auguste, RGD-Targeted Liposome Binding and Uptake on Breast Cancer Cells Is Dependent on Elastin Linker Secondary Structure. Bioconjugate Chemistry, 2016. 27(8): p. 1813-1821. Guo, P., et al., Using breast cancer cell CXCR4 surface expression to predict liposome binding and cytotoxicity. Biomaterials, 2012. 33(32): p. 8104-10. Panupinthu, N., H.Y. Lee, and G.B. Mills, Lysophosphatidic acid production and action: critical new players in breast cancer initiation and progression. Br J Cancer, 2010. 102(6): p. 941-6. Hama, K., et al., Lysophosphatidic Acid and Autotaxin Stimulate Cell Motility of Neoplastic and Non-neoplastic Cells through LPA1. Journal of Biological Chemistry, 2004. 279(17): p. 1763417639. Boucharaba, A., et al., The type 1 lysophosphatidic acid receptor is a target for therapy in bone metastases. Proceedings of the National Academy of Sciences, 2006. 103(25): p. 9643-9648. Chen, M., L.N. Towers, and K.L. O'Connor, LPA2 (EDG4) mediates Rho-dependent chemotaxis with lower efficacy than LPA1 (EDG2) in breast carcinoma cells. Am J Physiol Cell Physiol, 2007. 292(5): p. C1927-33. David, M., et al., Targeting lysophosphatidic acid receptor type 1 with Debio 0719 inhibits spontaneous metastasis dissemination of breast cancer cells independently of cell proliferation and angiogenesis. International Journal of Oncology, 2012. 40(4): p. 1133-1141.


42

Page 43 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72.

Mills, G.B. and W.H. Moolenaar, The emerging role of lysophosphatidic acid in cancer. Nat Rev Cancer, 2003. 3(8): p. 582-591. Liu, S., et al., Expression of Autotaxin and Lysophosphatidic Acid Receptors Increases Mammary Tumorigenesis, Invasion, and Metastases. Cancer Cell, 2009. 15(6): p. 539-550. Willier, S., E. Butt, and T.G. Grunewald, Lysophosphatidic acid (LPA) signalling in cell migration and cancer invasion: a focussed review and analysis of LPA receptor gene expression on the basis of more than 1700 cancer microarrays. Biol Cell, 2013. 105(8): p. 317-33. Sin, W.C., et al., G protein-coupled receptors GPR4 and TDAG8 are oncogenic and overexpressed in human cancers. Oncogene, 2004. 23(37): p. 6299-6303. Chen, P., et al., Gpr132 sensing of lactate mediates tumor–macrophage interplay to promote breast cancer metastasis. Proceedings of the National Academy of Sciences, 2017. 114(3): p. 580585. Murakami, N., et al., G2A Is a Proton-sensing G-protein-coupled Receptor Antagonized by Lysophosphatidylcholine. Journal of Biological Chemistry, 2004. 279(41): p. 42484-42491. Guo, Y., et al., Mannosylated lipid nano-emulsions loaded with lycorine-oleic acid ionic complex for tumor cell-specific delivery. Theranostics, 2012. 2(11): p. 1104-14. Son, C.H., et al., Combination treatment with decitabine and ionizing radiation enhances tumor cells susceptibility of T cells. Sci Rep, 2016. 6: p. 32470. Momparler, R.L., G.E. Rivard, and M. Gyger, Clinical trial on 5-aza-2'-deoxycytidine in patients with acute leukemia. Pharmacol Ther, 1985. 30(3): p. 277-86. Raychaudhuri, P. and H.J. Park, FoxM1: a Master Regulator of Tumor Metastasis. Cancer research, 2011. 71(13): p. 4329-4333. Jones, P.A. and P.W. Laird, Cancer epigenetics comes of age. Nat Genet, 1999. 21(2): p. 163-7. Yoo, C.B. and P.A. Jones, Epigenetic therapy of cancer: past, present and future. Nat Rev Drug Discov, 2006. 5(1): p. 37-50. Nervi, C., E. De Marinis, and G. Codacci-Pisanelli, Epigenetic treatment of solid tumours: a review of clinical trials. Clinical Epigenetics, 2015. 7: p. 127. de Lera, A.R. and A. Ganesan, Epigenetic polypharmacology: from combination therapy to multitargeted drugs. Clinical Epigenetics, 2016. 8(1): p. 105. Harris, J.M. and R.B. Chess, Effect of pegylation on pharmaceuticals. Nat Rev Drug Discov, 2003. 2(3): p. 214-221. Photos, P.J., et al., Polymer vesicles in vivo: correlations with PEG molecular weight. Journal of Controlled Release, 2003. 90(3): p. 323-334. Kubes, P. and C. Jenne, Immune Responses in the Liver. Annu Rev Immunol, 2018. 36: p. 247-277. Zhang, Y.N., et al., Nanoparticle-liver interactions: Cellular uptake and hepatobiliary elimination. J Control Release, 2016. 240: p. 332-348. Poelstra, K., J. Prakash, and L. Beljaars, Drug targeting to the diseased liver. J Control Release, 2012. 161(2): p. 188-97. Yung, Y.C., N.C. Stoddard, and J. Chun, LPA receptor signaling: pharmacology, physiology, and pathophysiology. J Lipid Res, 2014. 55(7): p. 1192-214. Lin, S., et al., Lysophosphatidic Acid Receptor 1 Is Important for Intestinal Epithelial Barrier Function and Susceptibility to Colitis. Am J Pathol, 2018. 188(2): p. 353-366. Keefe, D.M., et al., Chemotherapy for cancer causes apoptosis that precedes hypoplasia in crypts of the small intestine in humans. Gut, 2000. 47(5): p. 632-7. Jeynes, B.J. and G.G. Altmann, Light and scanning electron microscopic observations of the effects of sublethal doses of methotrexate on the rat small intestine. Anat Rec, 1978. 191(1): p. 1-17. Blanco, E., H. Shen, and M. Ferrari, Principles of nanoparticle design for overcoming biological barriers to drug delivery. Nature Biotechnology, 2015. 33: p. 941. Yao, C., et al., Highly Biocompatible Zwitterionic Phospholipids Coated Upconversion Nanoparticles for Efficient Bioimaging. Analytical Chemistry, 2014. 86(19): p. 9749-9757. Schmid, S.L. and L.L. Carter, ATP is required for receptor-mediated endocytosis in intact cells. J Cell Biol, 1990. 111(6 Pt 1): p. 2307-18.


43


73. 74. 75. 76. 77. 78. 79.

80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 93. 94.

Page 44 of 45

Batzri, S. and E.D. Korn, Interaction of phospholipid vesicles with cells. Endocytosis and fusion as alternate mechanisms for the uptake of lipid-soluble and water-soluble molecules. J Cell Biol, 1975. 66(3): p. 621-34. Choi, J.W., et al., LPA receptors: subtypes and biological actions. Annu Rev Pharmacol Toxicol, 2010. 50: p. 157-86. Yu, X., Y. Zhang, and H. Chen, LPA receptor 1 mediates LPA-induced ovarian cancer metastasis: an in vitro and in vivo study. BMC Cancer, 2016. 16(1): p. 846. Li, T.T., et al., Beta-arrestin/Ral signaling regulates lysophosphatidic acid-mediated migration and invasion of human breast tumor cells. Mol Cancer Res, 2009. 7(7): p. 1064-77. Gobeil, F., Jr., et al., Modulation of pro-inflammatory gene expression by nuclear lysophosphatidic acid receptor type-1. J Biol Chem, 2003. 278(40): p. 38875-83. Umezu-Goto, M., et al., Autotaxin has lysophospholipase D activity leading to tumor cell growth and motility by lysophosphatidic acid production. J Cell Biol, 2002. 158(2): p. 227-33. Liu, Z., et al., Characterization of decomposition products and preclinical and low dose clinical pharmacokinetics of decitabine (5-aza-2'-deoxycytidine) by a new liquid chromatography/tandem mass spectrometry quantification method. Rapid Commun Mass Spectrom, 2006. 20(7): p. 111726. Yoo, C.B., et al., Delivery of 5-aza-2'-deoxycytidine to cells using oligodeoxynucleotides. Cancer Res, 2007. 67(13): p. 6400-8. Lin, K.T., R.L. Momparler, and G.E. Rivard, High-performance liquid chromatographic analysis of chemical stability of 5-aza-2'-deoxycytidine. J Pharm Sci, 1981. 70(11): p. 1228-32. Raghavan, V., et al., Sustained Epigenetic Drug Delivery Depletes Cholesterol-Sphingomyelin Rafts from Resistant Breast Cancer Cells, Influencing Biophysical Characteristics of Membrane Lipids. Langmuir, 2015. 31(42): p. 11564-73. Vijayaraghavalu, S. and V. Labhasetwar, Efficacy of decitabine-loaded nanogels in overcoming cancer drug resistance is mediated via sustained DNA methyltransferase 1 (DNMT1) depletion. Cancer Letters. 331(1): p. 122-129. Berx, G. and F.V. Roy, The E-cadherin/catenin complex: an important gatekeeper in breast cancer tumorigenesis and malignant progression. Breast Cancer Research : BCR, 2001. 3(5): p. 289-293. Yoshiura, K., et al., Silencing of the E-cadherin invasion-suppressor gene by CpG methylation in human carcinomas. Proc Natl Acad Sci U S A, 1995. 92(16): p. 7416-9. Umbas, R., et al., Decreased E-cadherin expression is associated with poor prognosis in patients with prostate cancer. Cancer Res, 1994. 54. Siitonen, S.M., et al., Reduced E-Cadherin Expression is Associated With Invasiveness and Unfavorable Prognosis in Breast Cancer. American Journal of Clinical Pathology, 1996. 105(4): p. 394-402. Sulzer, M.A., et al., Reduced E-Cadherin Expression Is Associated with Increased Lymph Node Metastasis and Unfavorable Prognosis in Non-small Cell Lung Cancer. American Journal of Respiratory and Critical Care Medicine, 1998. 157(4): p. 1319-1323. Otto, T., et al., Inverse Relation of E-Cadherin and Autocrine Motility Factor Receptor Expression as a Prognostic Factor in Patients with Bladder Carcinomas. Cancer Research, 1994. 54(12): p. 31203123. Kiesslich, T., M. Pichler, and D. Neureiter, Epigenetic control of epithelial-mesenchymal-transition in human cancer. Molecular and Clinical Oncology, 2013. 1(1): p. 3-11. Rakha, E.A., et al., Prognostic markers in triple-negative breast cancer. Cancer, 2007. 109(1): p. 2532. Tang, D., et al., The expression and clinical significance of the androgen receptor and E-cadherin in triple-negative breast cancer. Medical Oncology, 2012. 29(2): p. 526-533. Zeng, Q., et al., CD146, an epithelial-mesenchymal transition inducer, is associated with triplenegative breast cancer. Proceedings of the National Academy of Sciences, 2012. 109(4): p. 11271132. Bektas, N., et al., Tight correlation between expression of the Forkhead transcription factor FOXM1 and HER2 in human breast cancer. BMC Cancer, 2008. 8: p. 42.


44

Page 45 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


95.

Yang, C., et al., FOXM1 promotes the epithelial to mesenchymal transition by stimulating the transcription of Slug in human breast cancer. Cancer Letters. 340(1): p. 104-112.


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