Poly(ethylene oxide monomethyl ether)-block-poly(propylene

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Poly(ethylene oxide monomethyl ether)-blockpoly(propylene succinate) nanoparticles. Synthesis and characterization, enzymatic and cellular degradation, micellar solubilization of paclitaxel and in vitro and in vivo evaluation Alessandro Jager, Eliézer Jäger, Zdenka Syrová, Tomaš Mazel, Lubomír Ková#ik, Ivan Raška, Anita Höcherl, Jan Ku#ka, Rafal Konefal, Jana Humajova, Pavla Pouckova, Petr Stepanek, and Martin Hruby Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b00048 • Publication Date (Web): 30 Mar 2018 Downloaded from http://pubs.acs.org on March 30, 2018

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Poly(ethylene oxide monomethyl ether)-blockpoly(propylene succinate) nanoparticles. Synthesis and characterization, enzymatic and cellular degradation, micellar solubilization of paclitaxel and in vitro and in vivo evaluation Alessandro Jäger†,‡,* Eliézer Jäger,†,*,‡ Zdenka Syrova,§,‡ Tomas Mazel,§ Lubomír Kováčik,ϕ Ivan Raška,§ Anita Höcherl,† Jan Kučka,† Rafal Konefal, †Jana Humajova,ɷ Pavla Poučková,ɷ Petr Štěpánek,† Martin Hrubý† †

Institute of Macromolecular Chemistry, Heyrovsky Sq. 2, 162 06 Prague, Czech Republic;

§

Institute of Biology and Medical Genetics, First Faculty of Medicine, Charles University and

General University Hospital in Prague, Albertov 4, 128 01 Prague, Czech Republic; ϕ

Institute of Cellular Biology and Pathology, First Faculty of Medicine, Charles University and

General University Hospital in Prague, Albertov 4, 128 01 Prague, Czech Republic; ɷ

Institute of Biophysics and Informatics, First Faculty of Medicine, Charles University in

Prague, Salmovska1, 120 00 Prague, Czech Republic;

ACS Paragon Plus Environment

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Keywords: polyester nanoparticles, enzymatic degradation, cellular uptake, paclitaxel, in vitro and in vivo cytotoxicity;

ABSTRACT: Polyester-based nanostructures are widely studied as drug delivery systems due to their biocompatibility and biodegradability. They have already reached the clinical use. In this work we describe a new and simple biodegradable and biocompatible system as the FDAapproved polyesters (PLGA, PCL and PLA) for the delivery of the anticancer drug paclitaxel (PTX) as a model drug. A hydrophobic polyester, poly(propylene succinate) (PPS) was prepared from a non-toxic alcohol (propylene glycol) and monomer from the Krebs’s cycle (succinic acid) in two steps via esterification and melt polycondensation. Further their amphiphilic block copolyester poly(ethylene oxide monomethyl ether)-block-poly(propylene succinate) (mPEO-bPPS) was prepared by three steps via esterification followed by melt polycondensation and the addition of mPEO to the PPS macromolecules. In vitro cellular behavior of the prepared NPs (enzymatic degradation, uptake, localization and FRET-pair degradation studies) were performed by fluorescence studies. PTX was loaded to the NPs of variable sizes (30, 70 and 150 nm) and their in vitro release was evaluated in different cell models being compared with commercial PTX formulations. The mPEO-b-PPS copolymer present Tg < body temperature < Tm, lower toxicity including the toxicity of their degradation products, drug solubilization efficacy, stability against spontaneous hydrolysis during transport in bloodstream and simultaneously enzymatic degradability after uptake into the cells. The detailed cytotoxicity in vitro and in vivo tumor efficacy studies have shown the superior efficacy of the NPs when comparable to PTX and PTX commercial formulations.


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INTRODUCTION A plethora of polymer delivery vehicles have been developed for targeted delivery of therapeutics to different tissues. Among the drug delivery systems, biodegradable and/or biocompatible polymer based nano- and microparticles have shown significant potential in selective delivery of diagnostics and therapeutics.1 The use of biodegradable/biocompatible polymers is very attractive because the controlled drug release can be optimized by suitable degradation strategies and it allows body clearance of the polymeric material avoiding its longterm accumulation and possible toxicity.2,3 Nevertheless, the available number of biodegradable polymers suitable for biomedical applications is limited due to the formation of harmful products and by-products from their degradation. So far, the use of biodegradable polymers aiming the preparation of nanoparticles carriers (NPs) and biomedical devices has been mostly limited to the aliphatic polyesters such as poly-ε-caprolactone (PCL), polylactic acid (PLA) and poly(lactic-coglycolic acid) (PLGA) essentially due their good hydrolizability, biocompatibility and drug release properties.1-4 From these polyesters several NPs encapsulating varied physicochemical different drugs, from hydrophilic to hydrophobic characteristics, were developed.5-8 Furthermore, due to the necessity of avoid rapid renal clearance and increase the stability of the drug-loaded NPs prepared from these polyesters, the production of PLGA, PLA and PCL-based blockcopolymers micelles from PLGA-b-PEO, PLA-b-PEO and PCL-b-PEO has also been reported3-8 as a strategy aiming the prolonged circulation of the NPs and their accumulation via the enhancement permeation and retention (EPR) effect.9-11 Through block copolymers self-assembly in aqueous media greater stability and drug targeting capabilities were achieved, as well as, decrease of dose-related side effects and increase of the solubility of hydrophobic molecules such as PTX.4,5,8,12 PTX, is a first-line treatment for


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breast, lung and ovarian cancer, and a second-line treatment for Kaposi sarcoma in HIV patients.4,13,14 However, because their poor water-solubility, PTX is usually formulated together with low molecular weight surfactants, e.g.Cremophor® EL (CrEL) and Tween 80® (polysorbate 80), which exhibit dose-related side-effects in humans. To overcome dose related side effects, PTX was formulated as NPs e.g., Abraxane®, one albumin bound PTX NPs formulation, that was the first NPs formulation approved product based on PTX.13,14 Preclinical and clinical studies demonstrated that Abraxane displays distinct pharmacokinetics and biodistribution properties, increased antitumor efficacy, and improved safety profile compared with CrEL-paclitaxel.14,15 As a result, Abraxane was approved for the treatment of multiple indications in chemotherapy.13-15 The clinical success of Abraxane demonstrates the great potential of NPs technology platforms such

as

PLGA,

PLA

and

PCL-based

block

copolymers

micelles

and

related

biodegradable/biocompatible systems. Taking into account the development of biomedical polymers candidates for the solubilization of PTX is a straightforward endeavor in the field of nanobiotechnology. Herein, we present the synthesis and characterization, the enzymatic and cellular degradation, the micellar solubilization of paclitaxel and the in vitro and in vivo evaluation of new NPs prepared from the poly(ethylene oxide monomethyl ether)-block-poly(propylene succinate) (mPEO-bPPS) block copolymer. When compared to the outstanding well established polyesters such, as PLGA, PLA or PCL the synthetized poly(propylene succinate) PPS copolymer shown similar physicochemical properties such as Tg < body temperature < Tm and lower toxicity including toxicity of degradation products. When NPs were prepared from the mPEO-b-PPS copolymer, drug solubilization efficacy and desirable release rates, stability against spontaneous hydrolysis during transport in bloodstream and simultaneously enzymatic degradability after uptake into the


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cells were achieved. Further detailed cytotoxicity in vitro and in vivo tumor efficacy studies shown the superior efficacy of the NPs when comparable to paclitaxel and paclitaxel formulations. All these features are presented along the manuscript.

EXPERIMENTAL SECTION Materials 1,3-Propanediol (PD), succinic acid (SA), poly(ethylene oxide) monomethyl ether (Mn ∼ 5,000 g⋅mol-1) (mPEO), titanium (IV) isopropoxide (TTiPO), Rhodamine B isothiocyanate and Reactive

blue

4

(1-amino-4[3-(4,6-dichlorotriazin-2-ylamino-2-ylamino)-4-

sulfophenylamino]anthraquinone-2-sulfonic acid) were purchased from Sigma-Aldrich (Czech Republic, CZ). Acetone, methanol, tetrahydrofuran (THF), dimethylformamide (DMF) and acetonitrile (ACN) were purchased from Merck (Czech Republic, CZ). Chromatographic grade THF was purchased from Fisher Scientific (Fisher Chemical, UK) and used for SEC analysis. All solvents, unless otherwise stated, were used without further purification. PTX was purchased from Santa Cruz Biotechnology (USA). Macrophages Raw 264.7, HeLa, MCF-7 and 4T1 tumor cell lines were obtained from American Type Cell Culture (ATCC) and cultured according ATCC guidelines. The water used was pretreated with the Milli-Q® Plus System (Millipore Corporation).

Methods PPS and mPEO-b-PPS synthesis. Poly(propylene succinate) (PPS) was synthesized following a previously described two-step melt polycondensation (esterification and polycondensation) protocol.16 Briefly, in a glass reactor were loaded the proper amount of SA


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and PD in an acid/diol molar ratio 1:1.2 and the catalyst TTiPO (4 × 10-4 mol⋅mol-1 diacid). The vessel was further evacuated and filled with argon. The reaction mixture was heated at 180 °C and stirred at constant speed (500 rpm). This first step (esterification) was considered complete when the theoretical H2O yield was removed from the reaction mixture by distillation and collected in a graduate cylinder (about ~ 3 h). The polycondensation reaction was carried out at 245 °C, 0.03 atm, under stirring at a constant speed (700 rpm) and it has been completed after 2 hours. The PPS polyester was purified via dissolution in chloroform and precipitation with methanol. In order to prepare the PEOylated PPS (mPEO-b-PPS) block copolymer a PPS with lower molecular weight was prepared by shortening the polycondensation time during the second step to 30 minutes. Then, a third step was employed according to Vassiliou et al.,16 during which 5 equiv. of mPEO of molecular weight 5000 g·mol-1 was added being further reacted at 240 °C for 30 min under vacuum (5.0 Pa). The mPEO-b-PPS was purified by repeated dissolution in chloroform and precipitation in cold methanol (3 times), dialyzed against water during three days (to remove unreacted mPEO) and recovered by lyophilization. Aminolysis of the PPS polyester (NH2-PPS). 1,6-Hexanediamine (1 Equivalent in DMF 25 mL) was dropwise added to a solution of the synthesized polyester PPS (1:2 Equivalent) in dried DMF (50 mL) over 10 minutes at room temperature. The temperature was raised up to 45 °C followed by stirring overnight. The polymer was recovered as NH2-PPS by lyophilization after being dialyzed for 48h against water. Synthesis of the Rhodamine B isothiocyanate-polyester conjugated (Rhoda-PPS). Rhodamine B isothiocyanate was added (1 mg·mL-1) to a solution of the synthesized polyester NH2-PPS in dried DMF (10 mL) and the solution was stirred for 24 h at 2-4 °C at dark. The obtained Rhoda-PPS conjugated was precipitated in cold methanol and dialyzed against


6

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deionized water for 5 days at room temperature and at dark. The deionized water was regularly exchanged every 12 hours to remove unreacted Rhodamine B isothiocyanate. The Rhoda-PPS conjugated was recovered after lyophilization. The Rhodamine B isothiocyanate concentration in the Rhoda-PPS conjugated was quantified by UV-vis. Synthesis of the Reactive Blue-polyester conjugated (Blue-PPS). Reactive Blue 4 (0.001 mol, 0.637 g) and triethylamine (0.003 mol, 0.303 g) were added to a solution of 20 mg·mL-1 of the synthesized NH2-PPS polyester in dried DMF (50 mL) at room temperature under stirring at dark. The temperature was raised up to 45 °C followed by stirring overnight. The polymer was recovered as PPS-Blue conjugated by lyophilization after being dialyzed for three days against water. Nanoparticles preparation. NPs were produced by nanoprecipitation technique.17 For the preparation of the NPs with different sizes typically, 12.5 mg of mPEO-b-PPS copolymer plus a certain amount of PPS copolymer (see Table 2) were dissolved in 5 mL of acetonitrile and injected using a syringe pump (EW-74900-00, Cole-Parmer®) into pure water or phosphate buffer saline solution (PBS) (10 mL) under stirring (1000 rpm). The organic solvent was further removed by evaporation under reduced pressure at room temperature and the aqueous solution was concentrated to the desired volume. NPs with different sizes loaded with certain amount of PTX (Table 2) were also produced in the same way as aforementioned (PTX was dissolved in the acetonitrile phase). The NPs containing rhodamine B with different sizes were prepared as follows: 1 mg of Rhoda-PPS copolymer conjugated was dissolved with 12.5 mg of the mPEO-b-PPS copolymer plus certain amount of PPS copolymer (Table 2) in acetonitrile (2.5 mL) following by injection of the organic phase using a syringe pump (EW-74900-00, Cole-Parmer®) into pure


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water or PBS (5.0 mL) under stirring (1000 rpm). The organic solvent was further evaporated as aforementioned. The NPs containing rhodamine B and Reactive-Blue (mix-NPs) were prepared as follows (Table S1): 0.3 milligrams of Rhoda-PPS copolymer conjugate was dissolved with 3 mg of the BluePPS copolymer conjugated plus 5 mg of the mPEO-b-PPS copolymer in acetonitrile (2.5 mL) following by injection of the organic phase into water or PBS (5.0 mL). The organic solvent was further evaporated as aforementioned. For the 1H NMR studies the NPs were prepared as previously, however, the injection of the organic phase was into deuterated water (D2O). Dynamic Light Scattering (DLS): DLS measurements were performed using an ALV/CGS-3 compact goniometer system consisting of a 22 mW HeNe linearly polarized laser operating at a wavelength of 632.8 nm, an ALV 7004 digital correlator and a pair of avalanche photodiodes operating in the pseudo cross-correlation mode. The samples were placed in 10 mm diameter glass cells and maintained at a constant temperature of 25 ± 1 °C. The autocorrelation functions reported were based on three independent runs of 60 s of counting time. The data were collected and further averaged by using the ALV Correlator Control software. The averaged intensity correlation functions g2 (t) were analyzed using the algorithm REPES18 (incorporated in the GENDIST program), resulting in distributions of relaxation times - A(τ ). The hydrodynamic radius (RH) of the nanoparticles was determined by using the Stokes-Einstein relation with D = τ -1 -2

q :

k BT q 2 RH = τ 6πη

(1)

where kB is the Boltzmann constant, T is the absolute temperature, q is the scattering vector,η is the viscosity of the solvent and τis the mean relaxation time related to the diffusion of the NPs.


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For this study, the distribution of relaxation times were also converted to distributions of RH by using the Stokes-Einstein equation. The polydispersity of the NPs was accessed by using the cumulant analysis18 of the correlation functions measured at 90o as: µ2 2 (2) t .... 2 where C is the amplitude of the correlation function and Γ is the relaxation frequency (τ −1). The ln g1 (t ) = ln C - Γ t +

parameter µ2 is known as the second-order cumulant and was used to compute the polydispersity index of the samples (PDI = µ 2 /Γ 2). Electrophoretic Light Scattering (ELS). ELS measurements were used to determine the average zeta potential (ζ) of the NPs, which were collected using a Zetasizer Nano-ZS ZEN3600 instrument (Malvern Instruments, UK). This instrument measures the electrophoretic mobility (UE) and converts the value to ζ-potential (mV) through Henry’s equation:

UE =

2 ε ξ f(ka) 3η

(3)

where ε is the dielectric constant of the medium and η is the viscosity. Furthermore, f(ka) is the Henry’s function, which was calculated through the Smoluchowski approximation f (ka) = 1.5. Each ζ-potential value reported in the manuscript was an average of 10 independent measurements with repeatability ± 2%. Small angle X-ray scattering (SAXS). The SAXS experiments were conducted at the high brilliance beamline ID02 of the European Synchrotron Radiation Facility (ESRF). The wavelength (λ) of the incoming beam and the sample-to-detector distance was chosen in such way that the targeted q range, being q = (4π/λ) sin(θ/2) (θ is the scattering angle) was covered. The collimated beam crossed sealed borosilicate capillaries (∼ 2 mm diameter) and was scattered to a X-ray image intensified low noise CCD (FReLoN) detector placed in an evacuated flight


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tube. The images were found to be isotropic, and the treatment has been made taking into account the 360° azimuthal scan. The resulting I(q) vs. q curves were corrected by the subtraction of the scattering water and they could be fitted by using spherical copolymer micelle model developed by Pedersen and Gerstenberg.19 The fitting procedures were performed using the SASfit software package developed by J. Kohlbrecher.20 Size exclusion chromatography (SEC) analysis. The weight-average molecular weight (Mw), number-average molecular weight (Mn) and the respective polydispersity I = (Mw/Mn) were obtained by SEC analysis. The SEC of the isolated polymers was performed at 25 °C with two PLgel MIXED-C columns (300 × 7.5 mm, SDV gel with particle size 5 µm; Polymer Laboratories, USA) and with UV (UVD 305; Watrex, Czech Republic) and RI (RI-101; Shodex, Japan) detectors. THF was used as a mobile phase at a flow rate of 1 mL⋅min-1. The molecular weight values were calculated using Clarity software (Dataapex, Czech Republic) using polystyrene as calibration standards. UV-Vis absorption spectroscopy. The UV-Vis spectra were measured in quartz cuvettes using an Evolution 220 UV-Visible Spectrophotometer (Thermo Scientific). The purified polyesters were measured using the scan mode (800 to 350 nm) at the scanning speed of 600 nm min-1 (0.5 nm steps). Calibration curves were prepared in DMF using Rhodamine B isothiocyanate and Reactive Blue at concentration range of 0.5 to 10 µg·mL-1 using the same above-mentioned setup. 1

H NMR measurements. 1H NMR spectrum of the polymer-conjugates and copolymers

were obtained using a Bruker Avance DPX 300 MHz NMR spectrometer or Bruker 600 MHz NMR spectrometer with CDCl3 as solvent (if not otherwise stated) at 25 °C and at 37 °C for polymer

degradation

studies.

The

chemical

shifts

are

relative

to

TMS

using


10

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hexamethyldisiloxane (HMDSO, 0.05pm from TMS) as internal standard. Chemical shifts as, δ, in units of parts per million (ppm). 1

H NMR measurements of the enzymatic NPs degradation. The mPEO-b-PPS NPs (1.0

mg·mL-1; D2O) were placed into a NMR tube and 1.1 U·mL-1 of Lipase in deuterated PBS buffer solution was also added. The 1H NMR degradation study (37°C) were recorded using a Bruker Avance III 600 spectrometer. T2 relaxation time values of PEO were determined from the linewidths using the relation T2 = (π∆ν)-1. Enzyme mediated NPs degradation studies. The degradation of the NPs was studied in enzymatic and non-enzymatic medium according previously published methodology.21-23 Briefly, the NPs (1.0 mg⋅mL-1) were loaded into 50 mL bottle (40 mL of mPEO-b-PPS NPs suspensions) and the volume was adjusted to 50 mL with phosphate buffer solution (PBS pH 7.4). To the bottles 1.1 U·mL-1 of Lipase were added and the bottles were sealed with parafilm and insulated from the air. Afterwards the bottles were incubated at 37 °C in dark. At predetermined intervals, samples were taken out from the bottles and lyophilized for SEC analysis (see Materials and Methods). Paclitaxel (PTX) drug loading and loading efficiency. The total amount of the chemotherapeutic PTX loaded into the NPs was measured by high performance liquid chromatography (HPLC, Shimadzu, Japan) using a reverse-phase column Chromolith Performance RP-18e (100 × 4.6 mm), eluent water-acetonitrile with acetonitrile gradient 0-100 vol%, flow rate = 1⋅0 mL⋅min-1, as described previously.24,25 The aliquote (100 µL) of drugloaded NPs was collected from the bulk sample and diluted to 900 µL with acetonitrile. Afterwards, 20 µL of the final sample was injected through a sample loop. PTX was detected at 230 nm using ultraviolet (UV) detection. The retention time of PTX was 3.0 min under such


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experimental conditions. An analytical curve with linear response in the range 0.5 – 100 µg⋅mL-1 was obtained and used to determine PTX contents (Figure S1). The drug-free was separated from the drug-loaded NPs by ultrafiltration-centrifugation (Ultrafree-MC 10,000 MW, Millipore) as previously detailed.26,27 The samples were centrifuged at 6000 rpm for 30 min. The free PTX amount was measured in the filtrated after the dissolution of NPs by using acetonitrile as described earlier. The drug-loading content (LC) and the drug-loading efficiency (LE) were calculated by using the following equations: ‫ ܥܮ‬ሺ%ሻ =

ௗ௥௨௚ ௔௠௢௨௡௧ ௜௡ ே௉௦

‫ ܧܮ‬ሺ%ሻ =

ௗ௥௨௚ ௔௠௢௨௡௧ ௜௡ ே௉௦

௠௔௦௦ ௢௙ ே௉௦

ௗ௥௨௚ ௙௘௘ௗ௜௡௚

‫ ݔ‬100 (4) ‫ ݔ‬100 (5)

Release experiments. The release experiments were carried out at 37 °C in pH adjusted release media (pH 7.4). Aliquots (500 µL) of PTX-NPs were loaded into 36 Slide-A-Lyzer MINI dialysis microtubes tubes with MWCO 10 kDa (Pierce, Rockford, IL). These microtubes were dialyzed against 4 L of PBS (pH 7.4) containing or not 1.1 U·mL-1 of Lipase. The release media was changed periodically to reduce the possibility of drug-diffusion equilibrium. The drug release experiments were done in triplicate. At each sampling time, three microtubes were removed from the dialysis system and 0.1 mL from each microtube was sampled and the remaining drug was extracted by using the aforementioned methodologies. The reported data are expressed as the amount of released PTX relative to the total PTX content in the PTX-NPs. Stability of the NPs in Simulated Physiological Media. The stability of the NPs was performed in 10 % blood plasma (Sigma-Aldrich) diluted in PBS (pH 7.4). Briefly, the NPs were incubated at 37 °C at concentration equal to 1 mg·mL-1. At predetermined intervals the DLS measurements were performed to probe the hydrodynamic radius and size-dispersity index of the entities in function of time.24


12

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Light microscopy. For live cell microscopy, HeLa and 4T1 cells were cultured in glass bottom Petri dishes (MatTek). Live cell imaging was performed using a spinning disc confocal system based on Olympus IX81 microscope equipped with Olympus UPlanSApo 100×/1.4NA oil immersion objective, CSU-X spinning disc module (Yokogawa) and Ixon Ultra EMCCD camera (Andor). Cells were maintained at 37°C and 5% CO2 with a microscope incubator (Okolab). All the NPs were optimized for containing 100 ng·mL-1of rhodamine B for the uptake experiments. Image analysis was performed in ImageJ and Matlab. For the in vitro cell degradation experiments, HeLa cells were incubated for several time points at 75 µg·mL-1 Rhodamine B. Data was acquired at an Olympus FV 10-ASV confocal microscope, 60x oil objective. Fluorescence was detected in Channel 1 (RhodB, exc. 543 nm, em. 625/50 nm) and Channel 2 (Reactive Blue, exc. 635 nm, em. 705/50 nm). In vitro Cytotoxicity. The MTT cytotoxicity assays were performed when cells became confluent. For the metabolic activity MTT assay, cells were cultured at a cell density of 5,000 for HeLa cells, 15,000 for Raw 264.7 cells, 10,000 for MCF-7 and 5,000 for 4T1 cells/well in a 96well microtiter plate. After 24 h of incubation at 37 ºC in 5% CO2, the cell culture medium was replaced by 100 µL of fresh culture medium containing tested NPs or free paclitaxel. After further 24 h of incubation at 37 ºC and 5% CO2, the medium was aspirated, and cells were incubated with 50 µL of MTT solution (1 mg·mL-1 in PBS) at 37 ºC in 5% CO2 for 2 h. Thereafter, MTT solution was aspirated and 100µL isopropanol was added. Synergy H1 Hybrid Reader instrument (BioTek, USA) and Sunrise (TECAN) microplate reader was used to assess cell viability by spectrophotometry at a wavelength of 570 nm (reference wavelength 690 nm). Results of MTT assay were expressed as percentage of controls (cells in control medium), which was considered as 100%. The tests were performed on at least three separate occasions.


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In vivo antitumor effect of the NPs. The animal experiments described in this manuscript were performed in accordance with the corresponding legislation in the Act on Experimental Work with Animals (Act No. 246/1992 of the Czech Republic and Decrees No. 419/2012), which is fully compatible with the corresponding European Union directives. For antitumor effect of the NPs, we used healthy inbred Balb/c mice (females, 8 weeks old, obtained from AnLab, Ltd., Prague, Czech Republic). After shaving the mice, 4T1 mammary carcinoma cells (1 × 106 cells in phosphate buffer) were injected subcutaneously into the right flank. The animals were randomly divided into three groups (n = 8). Subsequently, first group received NPs-Tx-formulation (60 mg PTX(equivalent)/kg) intravenously into the tail vein on day 4, 8 and 12 after injection of tumor cells. Second group received Taxol-Formulation (10 mg PTX(equivalent)/kg in the same way, but only on day 4 and 12 because of toxic symptom’s (skin irritation in the local of the injection) of the commercial formulation observed in the animals after the first dose. The last group served as untreated control, which received 200 µL of PBS on day 4, 8 and 12 after injection of tumor cells. Subsequently, survival of all animals was observed 60 days and tumor volume was measured twice a week (according to equation

௔×௕×௕×గ ଺

, where a

is the length and b is the width of tumor).

RESULTS AND DISCUSSION Synthesis and characterization of the copolyesters. The poly(propylene succinate) (PPS) copolymers were prepared by two step via esterification and melt polycondensation (step 1 and step 2, Scheme 1). After two hours of polycondensation reaction a high molar mass PPS was produced (Table 1) and subsequently used to conjugate the dyes (Scheme 2). The PEOylated counterpart poly(ethylene oxide) monomethyl ether-block-poly(propylene succinate) (mPEO-b-


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PPS) was prepared by three steps via esterification followed by melt polycondensation and by the addition of mPEO to the PPS macromolecules (step 3, Scheme 1).28 However, for this case the reaction time during the polycondensation (step 2, Scheme 1) was shortened to 30 minutes (instead of 2h) and therefore a PPS with lower molar mass was produced during the second step (Table 1). The resulted short PPS produced was directly reacted with mPEO during 30 minutes given the amphiphilic mPEO-b-PPS copolyester. The reaction between mPEO and the PPS end groups is either by an esterification or transesterification producing water or 1,3-propanediol as by-product (Scheme 1). Step 1 CH3

O HO

OH

Ti4+

OH

O

H

4

O

+

O

-

O

H3 C

+

H

O

O

180 °C

H 2O

O

OH

Step 2 CH3

O H3C

n

4

H

O

O

O

Ti4+

-

O

O

H

O

H

H

O

O

240 °C

O

O

+

n H 2O

n

Step 3 CH3

O H

O O

H

O O

n

+

H3 C

H3C

O O

-

4

H m

O

Ti4+

O

H

O O

240 °C

O O n

O m

CH3

+

H 2O

Scheme 1. Three step synthesis of the amphiphilic block copolyester poly(ethylene oxide) monomethyl ether-block-poly(propylene succinate) using titanium (IV) isopropoxide as catalyst.

In order to synthetize PPS polyester conjugates with Rhodamine B isothiocyanate and Reactive Blue, aminolysis of the carbonyl in the ester groups of PPS was performed using 1,6hexanediamine under mild conditions producing PPS amino terminated (NH2-PPS) according to the procedure adapted from literature29-31 (Scheme 2, step 1).


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Step 1 O

O

O O

HO

O

O

O O

+

OH

DMF

NH2

H2N

H2N

45 °C

O O

NH

n

O

O

NH O

O

NH2

n

Step 2 H3C H3C

N O

CH3 N CH3

OH

O

+

H2N

O O

O

NH

NH2

NH O

n

O

O

N C

CH3

S

H3C

CH3

DMF

N

2-4 °C

H3C N

O H3C H3C

O

N

S

O O

NH

NH

O

C NH

NH

O

O

N

S

O

HO

NH

nO

OH NH

CH3 CH3

O

Scheme 2. Two step synthesis of the Rhodamine B isothiocyanate-poly(propylene succinate) (Rhoda-PPS) under mild conditions.

Subsequently, NH2-PPS was reacted via nucleophilic addition to the NCS group32 of Rhodamine B isothiocyanate and via nucleophilic substitution to the chlorine of the triazine ring of the Reactive Blue 4 producing the Rhodamine B isothiocyanate-poly(propylene succinate) conjugate (Rhoda-PPS) and Reactive Blue 4-poly(propylene succinate) conjugate (Blue-PPS), respectively (showed for Rhodamine B, Scheme 2, step 2). The chemical structure of the prepared polymers was identified with

1

H NMR

spectroscopy and depicted in Figure 1, 2 and 3.


16

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Figure 1.1H NMR spectrum of PPS block (left) and mPEO-b-PPS diblock copolymer (right) in CDCl3.

Figure 2.1H NMR spectrum of Rhoda-PPS in CDCl3. Signals marked as “S” are related to THF.

Figure 3.1H NMR spectrum of Blue-PPS in CDCl3.


17


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As seen by 1H NMR (Figure 1, left), the PPS’s succinic acid methylene (b) protons appear as a singlet at δ = 2.62 ppm, while the propylene glycol methylene (a triplet, c pentet) protons are at δ = 4.16 and 1.96 ppm, respectively. The success of the coupling reaction of PPS with mPEO was confirmed by 1H NMR (Figure 1, right) and GPC data (Figure 4, discussed hereafter). The 1H NMR of the prepared mPEO-b-PPS show the typical peak at about δ = 3.65 ppm for the methylene protons of the mPEO block (b) and the singlet signal at δ = 3.39 attributed to the protons of methyl from the methoxy end group (a), which are absent in the spectrum of the prepared PPS. The signal of the protons corresponding to the succinic acid methylene protons appeared at δ = 2.60 ppm (c), while the propylene glycol methylene (a triplet,

c pentet) protons at δ = 4.14 and 1.94 ppm, respectively. The presence of methylene protons in purified copolymers and the absence of peaks related to free hydroxyl or carboxylic groups reveal that most of the PPS macromolecules have reacted with mPEO forming di- and triblock copolymers. The 1H NMR spectrum of Rhoda-PPS after nucleophilic addition of NH2-PPS to the NCS group of Rhodamine B isothiocyanate in order to produce the corresponding thiourea functional group (R1-NHCSNH-R2) is shown in Figure 2. Chemical structures and signal assignments are presented at the same figure. Although the high molar mass of the Rhoda-PPS difficult the identification of each peak position from the hydrogens of Rhodamine B in the 1H NMR spectra the presence of the regions containing the characteristics hydrogens from thioureahexeneamide at δ = 6.65 to 7.25 ppm (13), δ = 3.30 to 3.62 (14,19) and δ = 0.85 to 1.55 ppm (15,16,17,18) and xanthene ring at δ = 6.60 to 7.45 ppm (4,5,8,9), δ = 6.00 ppm (3), δ = 5.30 ppm (6) and δ = 4.55 ppm (7) with its respective hydrogens from the aromatic ring (10, 11, 12 at δ = 6.60 to 7.45 ppm


18

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Biomacromolecules

and diethylamine groups at δ = 3.30 to 3.62 (2) and δ = 0.85 to 1.55 ppm (1), confirms the covalent attachment of the dye to NH2-PPS polyester. Instead, PPS polyester shows typical strong signals from succinic acid methylene protons at δ = 2.60 ppm (20), while the propylene glycol methylene protons (21, 22), appears at δ = 4.14 and 1.94 ppm, respectively. The 1H NMR spectrum of Blue-PPS in CDCl3 with chemical structure and signal assignments is shown in Figure 3. Similarly to Rhoda-PPS the high molar mass of the Blue-PPS provide difficulties in the identification of each peaks position from the hydrogens of reactive Blue end groups which are just slightly visible. Signals of aromatic and NH protons (1, 2, 5, 6, 7, 8, 9) appears in δ = 8.4-6.8 ppm. Peaks related to Blue methyl groups (3) and methylene protons attached to nitrogen of hexanediamine (10, 15) are in the region of the spectrum with δ = 3.5-2.9 ppm. Signals of the rest of methylene protons of hexanediamine (11, 12, 13, 14) appears at δ = 1.6-1.2 ppm. Peaks of repeating units of polymer (16, 17, 18) are at δ = 2.60, 4.14 and 1.94 ppm, respectively. The SEC curves for the synthesized polyesters are shown in the Figure S2 (Supplementary Information file) and their physicochemical characteristics are shown in Table 1.

Table 1.Physico-chemical and thermal properties of the synthesized polyesters. Sample

Mn (g·mol-1)a

Mw (g·mol-1)a

Mw/Mna

Mn (g·mol-1)b

Tg(°C)c

Tm (°C) c

∆Hm c

PPS

21 100

37 900

1.79

—

-29

44

45

NH2-PPS

21 900

38 100

1.73

—

-29

44

40

Rhoda-PPS

23 100

39 000

1.69

—

-27

46

42

Blue-PPS

24 000

38 800

1.62

—

-29

45

35

PPSd

7800

11000

1.41

-29

44

43

mPEO-b-PPS

12 200

16 100

1.32

16 500

-41

50

63

mPEO

4500

5000

1.11

—

-53

59

205

a

b

1

c

d

measured by SEC. measured by H NMR. measured by DSC. short PPS, produced to react

with mPEO.


19


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The SEC curve for mPEO-b-PPS (Figure S2, right) shows a monomodal distribution with a symmetric peak which indicates absence of shoulders that appear at lower retention times due to unreacted mPEO chains (Figure S2, right). Thus, the purification process was successful in eliminate the unreacted mPEO chains and only di-and triblock copolymers are present in the purified sample. Indeed, the increase observed in the average molar mass of PPS after the mPEO reaction matches very well to the molar mass of mPEO-b-PPS with a diblock architecture (Table 1). Furthermore, the average number molar mass measured for mPEO-b-PPS by SEC and 1H NMR are in agreement if a diblock architecture for the copolymer is assumed which indicates that the reaction with mPEO takes place manly with the carboxylic end-group at one side of the polyester chain. This might be the case since the ratio of ~ 1:1 observed between the integrals from the signals of the carbons from the carboxyl and hydroxyl end groups in the

13

C NMR

spectra of PPS (Figure S3) as well as the concentration of carboxylic end-group measured ( ≈ 1.2

× 10-6 equivalents, data not shown) indicate the presence of about one group per chain. The SEC curves for the polyesters derived from the end group reactions with PPS (NH2-PPS, Rhoda-PPS and Blue-PPS) are shown in Figure S2 (left). The chromatograms show monomodal distribution for the polyesters after the end-group reactions with similar average molar mass distribution (Table 1) indicating that the reactions under mild conditions did not significantly affect their molar masses. DSC measurements were also employed in order to investigate how the reactions can affect the thermal behavior of synthetized polymers. The glass transition temperature (Tg), melting temperature (Tm) and heat of fusion (∆Hm) of the prepared polyesters and copolyester are presented in Table 1. Single Tg values between -29 to -27°C were observed for all polyesters after end group reactions (Figure S4, Supplementary Information file). Likewise, single values of


20

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Biomacromolecules

Tm ranging between 44 to 46 °C were also observed for the synthetized polyesters which are in agreement with previously published data16,33 and indicates that the end attachment did not changed their thermal properties. However, for mPEO-b-PPS copolymer a decrease on Tg and an increase on Tm values were observed in comparison to PPS homopolyesters indicating that the mPEO block affects the thermal properties of the prepared block copolyester. The presence of only one Tg value and its decrease in ~ 12 °C in comparison to the PPS homopolymers is a strong indication that some degree of miscibility between the mPEO and PPS blocks is present in the mPEO-b-PPS copolymer sample.16,34 It was previously described that PEOylation lowers the Tg values of mPEO-polyester diblock copolymers and its extension was related to the miscibility between the blocks in the amorphous state which is given by their molar mass and composition.34 Vassiliou et al. observed a decrease of about ~ 2 °C on the Tg of mPEO-b-PPS copolyester using mPEO of 2K at a molar composition of about ~ 37 mol%.16 Although only one Tg value was observed the authors were not convincing of miscibility between the blocks since the decrease observed on the glass transition was too low and close to the pure PPS. In our case, the mPEO 5K molar composition calculated by 1H NMR for mPEO-b-PPS was much higher (~ 60 mol%) and resulted in a greater decrease on the glass transition which indicates miscibility between the copolyester blocks in the amorphous state. Furthermore, the Tm recorded for mPEO-b-PPS was about ~ 5 °C higher in comparison to pure PPS (Table 1) and shows the influence of mPEO block melting temperature (~ 59 °C) on the Tm of the block copolymer. This was confirmed by the disappearance during the second heat of the shoulder attributed to the Tm of PPS block recorded on the thermogram at ~ 45 °C during the first heat of the mPEO-b-PPS (Figure S4). After the first cycle of heat and cooling the thermal history was erases and only a sharp melting


21


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peak at ~ 50 °C was observed on the mPEO-b-PPS sample which is attributed to the recrystallization of the mPEO block.34 The characteristic UV-Vis spectra of Rhoda-PPS and Blue-PPS polyester conjugates in DMF are shown in Figure S5. The amount of Rhodamine B isothiocyanate and Reactive Blue 4 covalently attached to Rhoda-PPS and Blue-PPS polyesters measured by the calibration curves in DMF were 10.4 and 15.2 µg·mg-1, respectively. As expected these values are smaller than the theoretical maximum amount calculated for Rhoda-PPS and Blue-PPS of 46.4 and 53.1 µg·mg-1, respectively, assuming a two end coupling of the amino polyester (NH2-PPS) with the probes. NPs preparation and characterization. The synthesized hydrophobic PPS polyester with high molar mass in combination with the amphiphilic mPEO-b-PPS block copolyester was employed in order to produce stable NPs with different sizes using the nanoprecipitation methodology. This methodology allows the polymers to fast nucleation and self-assembly after the contact of the organic with the aqueous phase ensuring the production of structurally defined NPs with a monodisperse size distribution.24,35,36 The expected assemblies are spherical coreshell NPs composed of a hydrophobic PPS core stabilized by the amphiphilic mPEO-b-PPS in which the mPEO blocks generates the sterically stabilized outer shell.37 The polymer concentrations used in the production of NPs and their resulted physicochemical characteristics determined by DLS/SLS and cryo-TEM are shown in Table 2.


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Table 2. Polymer concentrations in the organic phase and physicochemical properties of the produces NPs determined by DLS/SLS and Cryo-TEM. NPs sample

PPS (mg·mL-1)

mPEO-b-PPS (mg·mL-1)

Rhoda-PPS (mg·mL-1)

PTX (wt %)

DH (nm)

µ2/Γ2

mPEO-b-PPS30

0

5.0

0

0

29.3

0.18

mPEO-b-PPS70

5

2.5

0

0

72.5

0.15

mPEO-b-PPS150

10

2.5

0

0

157.8

0.11

mPEO-b-PPS-Rhoda30

0

5.0

0.3

0

28.2

0.12

mPEO-b-PPS-Rhoda70

5

2.5

0.3

0

68.9

0.15

mPEO-b-PPS-Rhoda150

10

2.5

0.3

0

164.2

0.09

mPEO-b-PPS-PTX30

0

5.0

0

5

32.1

0.15

mPEO-b-PPS-PTX70

5

2.5

0

5

78.2

0.08

mPEO-b-PPS-PTX150

10

2.5

0

5

160.6

0.13

By simple changing the amount of hydrophobic PPS in the organic phase was possible to fine tune the particles sizes without affecting their physiochemical properties. An example of the intensity size distribution measured by DLS and number size distribution measured by CryoTEM for the produced polymer NPs with different sizes are shown in Figure 4 and 5, respectively.

1.0

Normalized intensity, a.u.

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

Biomacromolecules

30 70 150

0.8

0.6

0.4

0.2

0.0 1

10

100

1000

Diameter, nm

Figure 4. Intensity-weighted size distribution of the mPEO-b-PPS-Rhoda NPs with different sizes in PBS pH 7.4 at concentration of 1 mg·mL-1.


23


(a)

(b)

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(c)

Figure 5. Cryo-TEM micrographs of polymer NPs with 30 nm diameter (mPEO-b-PPS30) in (a) (scale bar 20 nm); 70 nm NPs diameter (mPEO-b-PPS70) in (b) (scale bar 100 nm) and 150 nm diameter in (mPEO-b-PPS150) (c) (scale bar 200 nm).

The average intensity hydrodynamic diameter (2RH = DH) and polydispersity (µ2/Γ2) determined by DLS shows a monodisperse size distribution of mPEO-b-PPS-Rhoda NPs after preparation by the addition of different amounts of PPS polyester to the organic phase (Figure 4, Table 2). Accordingly, as PPS concentration increases in the organic phase from 0 to 10 mg·mL1

the NPs DH increases from 29.3 to 158 nm without significantly affecting the particles

dispersity (Table 2). In agreement with previously published data the trend was observed for all formulations and confirms the reliability and robustness of the method employed to prepare the NPs with different sizes.17 The fine tuning of the NPs size distribution was confirmed by CryoTEM measurements (Figure 5). Through the micrographs the diameters of the NPs were calculated from the average of at least 100 particles (see Materials and Methods). From the Figure 5a an average diameter of 18.5 nm (± 3.5 nm) were obtained, 66.5 nm (± 30.3 nm) for the Figure 5b and 98.1 nm (± 46.9 nm) for the Figure 5c. The experimental data suggest that the NPs


24

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diameters obtained from the cryo-TEM are in agreement with that determined by DLS. Moreover, the added concentration of PTX or Rhoda-PPS seems not to affect the size of mPEOb-PPS-PTX or mPEO-b-PPS-Rhoda NPs samples (Table 2). NPs enzymatic degradation. The enzymatic degradation of the prepared NPs was evaluated under the presence of the enzyme lipase from Pseudomonas sp. (1.1 U/mL)23 by DLS, SEC and 1H NMR. The NPs of different sizes were incubated with the predetermined concentration of the enzyme and the number average molar mass (Mn) was measured along the time of incubation (Figure 6). The decrease in the Mn of the NPs along the time of incubation with Lipase is evident (Figure 6a). Furthermore, a trend is observed between NPs degradation and NPs size being the smaller NPs degraded faster, as also evidenced after 48h (see SEC, Figure S6). Moreover, along the incubation time of the NPs with Lipase (1.1 U·mL-1) the overall scattering intensity was evaluated (see Methods) (Figure 6b). The NPs intensity remains mostly constant for the NPs of 70 and 150 nm as well as for the control NPs, NPs of 30 nm in PBS. However, for the NPs of 30 nm in presence of the Lipase the decrease of the overall scattering intensity is evident (Figure 6b, black circles). Taking into account that the intensity of scattering, Isc, is proportional to the concentration vs weight average molar mass according to the relationship, Isc ∼ cMw, the decrease in the overall scattering can be attributed to a decrease in either the particle’s molar mass (Mw) or polymer concentration (c).


25

Biomacromolecules

Average molar mass (g/mol)

12000

10000

8000

6000

4000

30 nm - Lipase 70 nm - Lipase 150 nm - Lipase 30 nm - PBS 70 nm - PBS 150 nm - PBS

2000

a)

b)

0 0

10

20

30

40

50

Degradation time (h)

OCH2 PPS C(O)CH2 PPS

0,9

OCH2 PEO

0,8

0,7

c)

Relaxation time T2 (ms)

85

1,0

I/I0

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

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80

75

70

d)

65

60

OCH2 PEO

0,6 0

10

20

30

40

50

60

70


0

10

20

30

40

50

60

70


Figure 6. Enzymatic degradation of mPEO-b-PPS NPs in presence of porcine lipase (1.1 U·mL1

) followed by SEC (a), DLS (b) and 1H NMR (c,d). The fastest degradation for the 30 nm NPs could be likely due to their smaller size and

consequently higher surface area available for Lipase to attack. Previous works using light scattering indicates that the Lipase biodegradation takes place with the enzyme attacking the NPs one by one, therefore, smaller NPs will be degraded much faster.38 The degradation of the mPEO-b-PPS NPs was also confirmed by 1H NMR measurements and by calculation of T2 relaxation times of PEO block from 1H NMR.39 The experimental data suggest that during the first hours the NPs are degraded primary at the ester bonds between the interface of PEO and PPS chains. This is evidenced by the increase on the mobility of mPEO chains observed by the T2 experiments (Figure 6d). At the same time decrease on the intensity of the signals from the polyester (Figure 6c) and the appearance of new signals from the degradation products were observed in the 1H NMR spectra (Figure S7). This process is quantitatively shown by the


26

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calculation of the ratio between Integral intensity “I” at the time “t” of the degradation and the Integral intensity from the starting time of experiment “I0” as depict in Figure 6c. The experimental data shown that in 10 hours ~ 20-30% of the polyester chains are degraded. Moreover no changes were observed along 72 hours from the NPs incubated at the same temperature without the presence of the enzyme (Figure S8). NPs drug release. To get more information’s related to the effect of the NPs sizes on the drug release, the in vitro drug release experiments were performed along 120h using a wellknown chemotherapeutic drug paclitaxel (see Methods for NPs preparation) and in the presence of Lipase (1.1 U·mL-1) (Figure 7). 30 nm - lipase 30 nm + lipase 70 nm - lipase 70 nm + lipase 150 nm - lipase 150 nm + lipase

100

Release of Paclitaxel, %

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

Biomacromolecules

80

60

40

20

0 0

20

40

60

80

100

120

time, hours

Figure 7. Drug release from Paclitaxel-loaded to mPEO-b-PPS NPs with 30 (black), 70 (blue) and 150 (red) nm diameter along time of incubation with 1.1 U⋅mL-1 of lipase (full circles) and PBS 7.4 (open circles) at 37 °C.

The release experiments at 37 °C were conducted also at pH 7.4 to simulate conditions during transport in blood and in normal healthy tissue. The results suggest that the drug release profile is clearly size and Lipase-sensitive. In general, for all the conditions investigated, the NPs releases the PTX chemotherapeutic more efficiently (around 1.5 times faster) in the presence of


27


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Lipase (Figure 7) when compared with the release in the absence of Lipase. For the smallest NPs with 30 nm the drug cargo is released almost twice as efficiently (∼92 % released) within 120 h in presence of Lipase (1.1 U·mL-1) than at physiological conditions of pH ∼ 7.4 and in absence of the enzyme. These results come in agreement with the NPs degradation experiments aforementioned. In vitro cytotoxicity effect of the mPEO-b-PPS NPs. The cytotoxicity studies of the mPEO-b-PPS-Rhod NPs, as well as, blank NPs (drug-unloaded mPEO-b-PPS NPs) was tested previously uptake studies in Hela cancer cell lines (Figure S9a) and Raw cells (Figure S9b). Different concentrations of the NPs (200 to 800 µg⋅mL-1) with the tree different sizes (30, 70 and 150 nm) were incubated along 24h for both cell lines. Cytotoxicity was not observed for blank NPs even at higher concentrations (800 µg⋅mL-1) in both of the cell lines. For mPEO-b-PPSRhoda NPs the viability was a little afected by the presence of the Rhodamine-B in Raw cells lines independent of NPs sizes or concentration, however, the viability values are still considered satisfactory and compared with values observed for the FDA-approved polymer poly-lactid acid (PLA).40 It is important to highlighted that Rhodamine B is considered toxic to cell lines41-43 at higher concentrations (Figure S9, RhodB) however, viability datas around 70% are considered normal especially at the higher concentrations of this study (800 µg·mL-1). Taking into account for the uptake studies the NPs concentration utilized were 8 times lower (~ 100 µg·mL-1). In vitro cellular uptake studies. The kinetics of the NPs cellular uptake was investigated using confocal light microscopy in HeLa and in 4T1 tumor cells lines. Live-cell imaging was employed using confocal spinning disc microscopy and the cells were seeded on Petri dishes with glass bottom (MatTek). Figure 8, 9 and S10 show the time course of cellular uptake and cellular distribution of the three tested NPs with different diameters (30 nm, 70 nm and 150 nm)


28

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Biomacromolecules

labeled with Rhodamine B isothiocyanate fluorescent dye in HeLa cells. The increase of fluorescence in cells was observed for all the three mPEO-b-PPS-Rhoda NPs with different sizes showing a distinct cellular distribution being followed. The fluorescent signal of 30 nm mPEO-bPPS-Rhoda NPs exhibited primarily vesicle-dependent accumulation in cytoplasm with mild diffuse cytoplasmic distribution in contrast to the 70 and 150 nm mPEO-b-PPS-Rhoda NPs that shown predominantly a dispersed cytoplasmic pattern. Nuclear localization of the NPs was observed in all cases. The time course of cellular uptake of mPEO-b-PPS-Rhoda NPs was plotted as a ratio of intracellular vs background fluorescence (Figure 9a). Data were fitted using a simple exponential to obtain the corresponding time constant (τ). The time courses of fluorescent signal and the corresponding time constant revealed clear differences between the tested NPs. The fluorescent signal increased for all NPs samples tested during 10-16 h however with distinct patterns. The cells incubated with NPs of 30 nm in diameter exhibited higher intracellular fluorescence in contrast to the cells incubated with 70 and 150 nm NPs, respectively. The lowest signal is demonstrated for 150 nm mPEO-b-PPS-Rhoda NPs (Figure 9a). Time course of cellular uptake of mPEO-b-PPS-Rhoda NPs and the corresponding time constant, τ, revealed that the 30 nm NPs are the NPs with the fastest increase of fluorescence τ = 278 min, followed by 70 nm NPs with τ = 455 min and 150 nm NPs τ = 627 min as the NPs with slower cell uptake (Figure 9b). Faster cellular uptake can result in more efficient cellular delivery therefore resulting in higher effects of payload drug in cells.


29


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Figure 8. Time course of cellular uptake of mPEO-b-PPS-Rhoda NPs. Increase in fluorescence after the addition of NPs of three different sizes (30, 70 and 150 nm) was observed in HeLa cells. Data are presented as a ratio of intracellular vs background fluorescence. The time courses were fitted using a simple exponential and the corresponding time constant (τ) was obtained.


30

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Figure 9: Time course of fluorescence increase in HeLa cells after the addition of mPEO-b-PPSRhoda NPs of three different sizes. A: Time course of the ratio of intracellular vs background fluorescence (black – 30 nm, white – 70 nm, gray – 150 nm NPs; mean ± SEM). B: Time course curves were fitted using a simple exponential and the corresponding time constant (τ) was obtained. Data presented as mean ± SEM (N=23, 16 and 11 for 30, 70 and 150 nm NPs).

The uptake studies on 4T1 mammary carcinoma cells lines showed a similar dependence of the particles sizes on the uptake of the mPEO-b-PPS-Rhoda NPs (Figure 10 and S11). It was observed that smaller the particles size are, larger was the increase on the fluorescence intensity during the time and therefore faster the particles uptake. The uptake is better visualized in Figure S11 were is depicted the time course of the ratio of intracellular vs background fluorescence for the tree different NPs sizes evidencing always the higher uptake along time for the smallest NPs, 30 nm as well as for the 4T1 cells. It is well known that endocytosis is the mechanism that governs NPs internalization and uptake by cells being dependent on the structural features and chemical characteristics of the assemblies because they influence the adhesion of the NPs and the “wrapping time” which described how a membrane enclosed a particle.44 In this way, NPs size, shape and charge have


31


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been already reported to have considerable implications in cell uptaken.44-46 Positively charged NPs are endocytosed to a greater extent when compared to neutral or negatively charged counterparts as well as related to the NPs shape, elongated assemblies are usually internalized less efficiently than spherical NPs as the result of longer membrane wrapping time required for elongated NPs.47,48 For spherical NPs the endocytosis mediated by clathrin or caveolin are the main pathways been suggested towards the internalization of sub-100 nm particles with minimum and optimum sizes of ~ 50 nm.49

Figure 10: Time course of cellular uptake of mPEO-b-PPS-Rhoda NPs. Increase in fluorescence after the addition of NPs of three different sizes (30, 70 and 150 nm) was observed in 4T1 cells.


32

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Biomacromolecules

Data are presented as a ratio of intracellular vs background fluorescence. The time courses were fitted using an exponential increase function and the corresponding time constant (τ) was obtained.

Independent of the cell line our uptake study indicates that smaller particles of ~ 30 nm sizes can cross the cell membrane and enter the cell much faster in comparison to bigger particles. Taking into account that the NPs surface chemistry is similar (covered by mPEO) and they are spherical in shape and relatively neutral in charge (data not showed) the differences founded in the uptake are only likely due to the different size of the NPs. In vitro NPs degradation experiments in living cells. Because of the biological relevance of the NPs with 30 nm i.e., presenting the higher uptake in cells and demonstrating the best in vitro performance, they were chosen for all the further experiments. NPs cellular degradation is important for both, payload release and particles elimination. For these we perform the experiment of NPs degradation (see Methods) based on quenching of the fluorescence of Rhodamine B (maximum emission wavelength of 580 nm) by Reactive Blue 4 (maximum absorption wavelength at 598 nm), both covalently attached to the NPs (named mix-NPs, see Methods). To have a satisfactory quenching of fluorescence different ratios of Rhoda-PPS and Blue-PPS copolymer conjugates were evaluated by fluorescence spectroscopy in PBS (Figure S12). Is worth to mention here that to find the best quenching ratio for the mix-NPs increasing amounts of Blue-PPS were added to the organic phase resulting in an increase in particles size. To compensate the increase on particles size caused by the addition of Blue-PPS larger amounts of mPEO-b-PPS were added to the organic phase in order to maintain the desired size (DH ~ 30 nm) of the mix-NPs. Consequently, to keep the same amount of polymer for degradation in both NPs systems the final polymer concentration in the mPEO-b-PPS-Rhoda NPs was corrected in


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relation to the mix-NPs by the addition of mPEO-b-PPS according to Table S1. The best quenching ratio measured by fluorescence spectroscopy between Rhoda-PPS and Blue-PPS in the mix-NPs within the desired size was 1:10 (Figure S12, Table S1). At this ratio a complete quenching of Rhoda-PPS was observed in the mix-NPs in comparison to the non-quenching mPEO-b-PPS-Rhoda NPs using the same Rhodamine concentration (Table S1). Furthermore, according to DLS (Figure S13) the average intensity hydrodynamic diameters and polydispersity’s show a monodisperse size distribution for both mPEO-b-PPS-Rhoda and mixNPs with similar sizes of around DH ~ 32 nm (Table S1 and Figure S13). These results indicate that the fluorescent mPEO-b-PPS-Rhoda NPs and the fluorescent quenched mix-NPs within the desired sizes were successfully prepared using the nanoprecipitation method. Their sizes within the targeted region of about DH ~ 30 nm and their specifically fluorescent properties allow them to be employed as standard NPs for the in vitro degradation studies and their behavior can be therefore extrapolated to help on the interpretation of other mPEO-b-PPS NPs systems such as those containing PTX which were employed in the cytotoxicity experiments in vitro and in vivo. Before the degradation experiments the neglected cell toxicity of the mix NPs was demonstrated in HeLa cells along 48h of incubation at different NPs concentrations (Figure S14). The time course of NPs degradation in HeLa cells is portrayed in the Figure 11. Is clearly evidenced that the onset of NPs degradation occurs after 72h of incubation, visible by the initially highly quenched RhodB fluorescence in the mix-NPs (Figure 11, middle row) and the following un-quenching causing a strong increase of the RhodB fluorescence from the mix-NPs at 72h incubation (Figure 11, middle row).


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Oppositely, the fluorescence from the NPs prepared just with RhodB where the Rhodamine fluorescence is not quenched by a second dye, is observed along all the times of incubation (Figure 11, top row). In these NPs prepared with only RhodB and without the quenching second dye, the RhodB signal observed already at begin of the incubation is stronger. These results indicated that the NPs are likely stable at least along 48h under the cell culture medium tested, a considerable advantage for desirable long circulation times of the NPs for further accumulation of the drug delivery system in the active targeted tumor sites.

0.5 h

4h

24 h

72 h

Figure 11: Time course of NPs degradation in HeLa cells lines from 0.5 h to 72h.


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In vitro cytotoxic studies of PTX-loaded and unloaded NPs. In order to verify if the dependence of the particles sizes on the uptake will exert differences on the cell cytotoxicity, the NPs with the different sizes were loaded with the anticancer drug PTX (Table 2) and the cellular cytotoxicity was tested in vitro against HeLa cancer cells and compared to PTX-unloaded NPs (Figure 12). The results have shown an important increase in the cytotoxicity of the PTX-loaded NPs in comparison with the free PTX (Figure 12a). The higher cytotoxicity of the NPs is expected because free PTX is bound to serum albumin in blood plasma in which form it is slowly uptaken by the cells.50 IC50 of tested 30 nm PTX-loaded NPs exhibited significant higher toxicity 8.0 +/- 1.3 nM in contrast to 70 nm PTX-loaded NPs 11.1 +/- 1.1 nM and 150 nm PTXloaded NPs 11.5 +/- 0.8 nM, being all of them more toxic compare to PTX 29.2 +/- 0.3 nM (Figure 12b). The noticeable reduction in the cell viability for PTX-loaded NPs in comparison to free PTX indicates a faster and more efficient uptake of the drug-loaded NPs in comparison to free drug. Furthermore, the dependence of the particles size on cytotoxicity was confirmed in agreement with the in vitro cell uptake experiments (Figure 8, 9).

Figure 12: Size dependent cytotoxicity of the Blank and PTX-loaded NPs with different sizes in HeLa cancer cell line along 24h of incubation. (A) Cell viability and the dose response of tested NPs. (B) The comparison of IC50 of PTX-loaded NPs 30 nm,70 nm and 150 nm, p ˂ 0,05.


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The negligible cytotoxicity observed for the PTX-unloaded NPs also evidenced that the cytotoxic effect is due to the release of the PTX from the NPs. These findings confirm the importance of the particles size on the cell uptake and cytotoxicity which directly influences in the results of the cancer therapy. Besides particles size and stability the preparation of NPs with a high drug loading content (eq. 4) is also one of the key relevant factors on the therapeutic efficacy of any nanomedicine formulation intended for application in cancer therapy.51 Taking it into account the authors tested the preparation of NPs of ~ 30 nm by loading the highest amount of PTX possible at the organic phase, however, ensuring the production of stable NPs without the presence of precipitates after the evaporation of the organic solvent. Thus, crescent amounts of PTX were added to the organic phase containing mPEO-b-PPS according to Table 3. Stable NPs were prepared up to a loading of 15 wt% of PTX without the presence of precipitates. The NPs were characterized by DLS and SAXS. As general, both scattering techniques shows a significate increase in the particles size after the addition of a threshold amount of PTX to the prepared NPs. However, unlike SAXS the DLS measurements show an increase in the NPs size only after addition of PTX at concentrations higher than 7.5 wt % (Table 3). This is explained taking into account the differences on the principles used for measure the particles size between both techniques. While DLS uses the fluctuations of the intensity of the scattered light of an object to calculate its diffusive size, assuming that it behaves as a homogeneous sphere when diffusing into a media, SAXS technique uses the differences in the scattering contrast (electron density) between the particles and the media allowing to the distinction of structural inner features of the NPs like e.g. core and shell.52 Thus, while hydrodynamic dimensions (DH) of the NPs were not affected by the loading of PTX at concentrations up to 7.5w t% the fitting of SAXS curves using


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the model of spherical copolymer micelle (Figure S15) shows that the radius of the particles core (Rcore) start to increase after the loading of PTX at 7.5 wt % (Table 3). The increase on NPs size with the increase on drug loading was observed by SAXS measurements only in the particles core. As previously observed this is a strong indication of a favorable polymer-drug interaction between the PPS core and PTX. 28,53,54 This is confirmed by the high amounts of PTX loaded to mPEO-b-PPS NPs in comparison to similar NPs systems commonly used as nanocarriers for drug delivery applications that uses similar polyester cores such as PLA, PLGA and PCL.55

Table 3. Polymer and PTX concentrations and the resulted sizes of the PTX loaded NPs measured by DLS and SAXS.

NP Sample

mPEO-b-PPS (mg.mL-1)

PTX (mg·mL-1)

PTX loading (wt%)

DH (nm)

µ2/Γ2

Rcorea (nm)

Rshella (nm)

mPEO-b-PPS

5

0

0

29.3

0.18

11

5.1

mPEO-b-PPS-PTX5

5

1

5

32.1

0.15

10.8

5.4

mPEO-b-PPS-PTX7.5

5

1.5

7.5

31.3

0.15

12.1

7.5

mPEO-b-PPS-PTX10

5

2

10

39.3

0.19

14.1

6.2

mPEO-b-PPS-PTX15

5

3

15

44.5

0.21

14.8

6.5

a

fitted by using the model of sphere with Gaussian chains attached.

As we target stable NPs with sizes of ~ 30 nm and loading higher amounts of PTX for further experiments the authors choose the mPEO-b-PPS-PTX10 NPs.

In vitro cytotoxicity studies of PTX-loaded NPs in comparison with commercial formulations. The authors performed further cytotoxicity experiments with the PTX-loaded NPs of 30 nm


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(mPEO-b-PPS-PTX5, Table 3) (named NPs-Tx) and compared then with a PTX commercial formulation (Mylan®, Paclitaxel injection, USP, 6 mg·mL-1; Czech Republic) (named TaxolFormulation), as well as, with a PTX solution (in 2% of DMSO)25,56 (named Tx) and with the NPs with the higher PTX encapsulation aforementioned (PEO-b-PPS-PTX10, Table 3), and named as NPs-Tx-formulation. The formulations were tested in 3 different tumor cell lines being two of them of breast cancer, MCF-7 and mouse 4T1, and one from cervical adenocarcinoma, HeLa, as well as in a non-tumor cell model, Abelson murine leukemia virus-induced tumor; ascites Raw 264.7 cells (Figure S16). The NPs-Tx-formulation for both of the tested breast cancer cell line exhibited significant lower IC50 values when comparing to free Tx. For MCF-7 cell line, the values decreases from 35.0 +/- 4.6 nM for free PTX to 10.2 +/- 0.8 nM for NPs-Tx-formulation and for 4T1 cell line the values decreases from 117.1 +/- 17.8 nM to 23.1 +/- 9.0 nM (Figure 13). There was a clear difference of IC50 values between the tumor cell lines, the 4T1 cell line was less sensitive to Tx ( ≥ three-times), however, in comparison of cytotoxicity free Tx/Tx from NPs-Tx-formulation, the NPs-Tx-formulation was more toxic in contrast to MCF-7 cells. Hela cell line exhibited the highest sensitivity to PTX; IC50 values of 29.2 +/- 0.3 nM and 7.8 +/- 0.5 nM for NPs-Txformulation in contrast to macrophages Raw 246.7 cell line. Taxol-Formulation, as the commercial drug, revealed higher cytotoxic effect to all of tested cell lines compare to Tx. The higher toxicity is related to the components of the formulation i.e., PEOylated castor bean oil and ethanol, and is a well-know and reported effect of these Tx formulation coming from membrane damage caused by these excipients.5,13,14


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Figure 13: IC50 of the NPs-Tx in cancer cell lines Hela, MCF-7, 4T1 and murine macrophages Raw 264.7 along 24h incubation in comparison with NPs-Tx-formulation, Taxol-Formulation and Tx.

Therefore taking into account, the higher cells sensitivity to PTX from the NPs-Tx and NPs-Tx-formulation (Figure S16 and 13), their higher kinetic of cell uptake (NPs of ~ 30 nm, Figure 8 and 9) and their faster PTX release (Figure 7); the higher cytostatic effect observed in the in vitro tests demonstrated how the ~ 30 nm NPs loading with PTX positively impacts on the efficiency for treating cancer cells. In vivo antitumor effect of the NPs. Because the positive results from the NPs-Txformulation in different cell lines we further decided to evaluate the antitumor efficacy of the NPs in vivo. Prior to in vivo studies the stability of the prepared NPs was tested in vitro simulated complex biological media (Figure S17). The negligible changes of the NPs intensity size distribution (Figure S17, left) and scattering intensity (Figure S17, right) in function of time for the NPs-Tx-formulation during incubation with diluted human plasma (10% in PBS along 24h) suggest that the NPs are highly stable against aggregation in the simulated physiological


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media.24,57 The slight increase in the NPs sizes are well documented (NPs sizes increases ~ 5 – 10 nm) and are related to the adsorption of a protein monolayer because the average size of the dissolved single proteins is ~8 nm.24,52 The anticancer activity of the NPs-Tx-formulation was evaluated in 4T1 tumor-bearing Balb/c mice compared with the Taxol-Formulation (Mylan® paclitaxel commercial formulation) and control group with PBS. The developed tumors killed the untreated mice within 34-51 days (the mean survival time was 43.3 days, SD = 6.28, median survival = 45 days). The TaxolFormulation administrated in two doses (2×10 mg PTX(equivalent)/kg) on the 4th and 12th days allowed survival within 37-47 days (the mean survival time was 43.7 days, SD = 3.56, median survival = 45 days). Because the negligible toxic effects of the NPs-Tx-formulation tested in healthy mice along the tests of NPs maximum tolerated doses the NPs-Tx-formulation was administered in tree doses (3×60 mg PTX(equivalent)/kg) on the 4th, 8th and 12th days after tumor transplantation. The developed tumors killed the mice treated with the NPs-Tx-formulation within 48-52 days (the mean survival time was 51.4 days, SD = 2.07, median survival = 52 days). The NPs-Tx-formulation was clearly more effective than the Taxol-Formulation as well as the control group in prolong the animal survivor (Figure 14). In comparison with controls, the average survival was prolonged about 8 days (P = 0.01778 and F = 7.53267 at α level 0.05). After 30 days of treatment the tumor growth was reduced by ~ 26 % and ~ 33,3 % when compared with the controls (P = 0.02905 and F = 6.1422 at α level 0.05) and with the TaxolFormulation (P = 0.04213 and F = 5.1715 at α level 0.05), respectively (Figure 14, left). The reason, why tumor volumes and survival of the group with Taxol-Formulation were comparable with the control group, can be explained by toxicity of formulation followed by dose reduction.


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Control (PBS) Taxol-Formulation NPs-Tx-formulation (60 mg/kg)

3.6

100

3.0

Surviving mice (%)

Tumor volume, cm3

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

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2.4

1.8

1.2

Controls Taxol-Formulation NPs-Tx-formulation 60 mg/Kg

80

60

40

20 0.6

0 0

4

8

12

16

20

24

28

32

0

10

Days after treatment

20

30

40

50

60

Time after treatment (days)

Figure 14: In vivo effect on the growth of 4T1 cell mammary carcinoma (left) and Kaplan-Meier survival plot of mice (right) of NPs-Tx-formulation, Taxol-Formulation and control (treated mice PBS).

Along with the reduction in tumor growth, the survival time of the animals was also extended over that of the untreated controls and the Taxol-Formulation by using the NPs-Tx-formulation (Figure 14, left).

CONCLUSIONS We describe a simple new biodegradable and biocompatible system as alternative to the FDAapproved polyesters for the delivery of the anticancer drug paclitaxel as a model drug. A hydrophobic polyester, poly(propylene succinate) (PPS) was successfully prepared from a nontoxic alcohol and a monomer from the Krebs’s cycle via esterification and melt polycondensation. Further their amphiphilic block copolyester (mPEO-b-PPS) was prepared via esterification followed by melt polycondensation and the addition of mPEO to the PPS macromolecules. In vitro cellular behavior of the prepared NPs (enzymatic degradation, uptake, localization and FRET-pair degradation studies) demonstrate simultaneously size dependent


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enzymatic degradability and higher uptake into different tumor cells. PTX when loaded to the NPs of variable sizes (30, 70 and 150 nm) shown size dependent in vitro release and cytotoxicity in different cell models (Raw 264.7, HeLa, MCF-7 and 4T1 tumor cells) outstanding commercial paclitaxel formulations. From selected 30 nm NPs more detailed cytotoxicity in vitro and in vivo tumor efficacy studies demonstrate the superior efficacy of the NPs when comparable to PTX and PTX formulations. The PTX- loaded mPEO-b-PPS NPs shown to be a promising drug delivery system for the chemotherapy.

ASSOCIATED CONTENT

Supporting information HPLC calibration curve for PTX, DSC scans of the synthesized copolyesters, UV-Vis absorption spectra of fluorescent copolymer conjugates, DLS of fluorescent NPs and from the stability of NPs in plasma, SEC chromatograms of NPs after enzymatic degradation, 1H NMR spectrum of the NPs after and along enzymatic degradation, uptake studies of the NPs in HeLa cells, SAXS patterns of PTX-loaded NPs. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION

Corresponding Authors *(A.J.) E-mail: [email protected]; *(E.J.) E-mail: [email protected];


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Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ‡These authors contributed equally.

Notes The authors declare no competing financial interest. ACKNOWLEDGMENT E. Jäger, A. Jäger, M. Hruby, J. Humajova and P. Poučkova acknowledge the Czech Science Foundation (grant #17-09998S). P. Štěpánek, I. Raška, Z. Syrova acknowledge the Czech Science Foundation (grant #17-07164S). The authors thanks to the Department of Biochemistry, Faculty of Science – Chemistry Section, Charles University and Dr. Olga Janoušková from Institute of Macromolecular Chemistry AS CR for their kind help with the instrumentation necessary for biological experiments.

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TOC - Graphical Abstract (For Table of Contents Use Only)

The superior in vivo suppression of tumor growth caused by paclitaxel (PTX) loaded mPEO-bPPS NPs in comparison to the commercial formulation Mylan® makes this new PEOylated polyester a promising drug delivery tool for the therapy of cancer.


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Poly(ethylene oxide monomethyl ether)-block-poly(propylene

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