Reductively Degradable Poly(2-hydroxyethyl methacrylate) Hydrogels

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Reductively degradable poly(2-hydroxyethyl methacrylate) hydrogels with oriented porosity for tissue engineering applications Hana Mackova, Zdenek Plichta, Helena Hlidkova, Ondrej Sedlacek, Rafal Konefal, Zhansaya Sadakbayeva, Miroslava Duskova-Smrckova, Daniel Horák, and Sarka Kubinova ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b01513 • Publication Date (Web): 13 Mar 2017 Downloaded from http://pubs.acs.org on March 18, 2017

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ACS Applied Materials & Interfaces

Reductively degradable poly(2-hydroxyethyl methacrylate) hydrogels with oriented porosity for tissue engineering applications Hana Macková*,a, Zdeněk Plichtaa, Helena Hlídkováa, Ondřej Sedláčeka, Rafal Konefala, Zhansaya Sadakbayevaa, Miroslava Dušková-Smrčkováa, Daniel Horáka, Šárka Kubinováb

a

Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic,

Heyrovsky Sq. 2, 162 06 Prague 6, Czech Republic b

Institute of Experimental Medicine, Academy of Sciences of the Czech Republic, Vídeňská

1083, 142 20 Prague 4, Czech Republic KEYWORDS: Poly(2-hydroxyethyl methacrylate), reductively degradable, oriented porosity, 2methacryloyloxyethyl phosphorylcholine, 2-(acethylthio)ethyl methacrylate, hydrogel

ABSTRACT

Degradable poly(2-hydroxyethyl methacrylate) hydrogels were prepared from a linear copolymer (Mw = 49 kDa) of 2-hydroxyethyl methacrylate (HEMA), 2-(acethylthio)ethyl methacrylate 1 ACS Paragon Plus Environment


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(ATEMA),

and

zwitterionic

2-methacryloyloxyethyl

phosphorylcholine

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(MPC).

The

deprotection of ATEMA thiol groups by triethylamine followed by their gentle oxidation with 2,2'-dithiodipyridine resulted in the formation of reductively degradable polymers with disulfide bridges. Finally, a hydrogel 3D structure with an oriented porosity was obtained by gelation of the polymer in the presence of needle-like sodium acetate crystals. The pore diameter and porosity of resulting poly(2-hydroxyethyl methacrylate-co-2-(acethylthio)ethyl methacrylate-co2-methacryloyloxyethyl phosphoryl-choline) [P(HEMA-ATEMA-MPC)] hydrogels varied between 59-65 µm and 70-79.6 vol.% according Hg porosimetry and complete degradation of these materials was reached in 86 days in 0.33 mmol solution of L-cysteine/l in phosphate buffer. The crosslinked P(HEMA-ATEMA-MPC) hydrogels were evaluated as a possible support for human mesenchymal stem cells (MSCs). No cytotoxicity was found for the uncrosslinked thiolcontaining and protected P(HEMA-ATEMA-MPC) chains up to a concentration of 5 and 1 wt.% in alpha-minimum essential medium, respectively.

INTRODUCTION Poly(2-hydroxyethyl methacrylate) (PHEMA), which have been used as implants1 due to their non-toxicity, inertness, and resemblance to living tissue2, it is generally non-degradable and insoluble in water. Moreover, 2-hydroxyethyl methacrylate (HEMA) contains residual impurities of cross-linker e.g., ethylene dimethacrylate, which makes the polymerized material insoluble in solvents. Many attempts to prepare degradable PHEMA hydrogels have been described. For example, the solubility of PHEMA in water was enhanced by the copolymerization of HEMA with hydrophilic methacrylates e. g. methacrylic acid3, [2-(methacryloyloxy)ethyl]trimethyl 2 ACS Paragon Plus Environment

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ammonium chloride3, 2-(N-3-sulfopropyl-N,N-dimethyl ammonium)ethyl methacrylate4, (2-O(N-acetyl-β-D-glucosamine)ethyl methacrylate5, itaconic acid6. An increased solubility in biological fluids was also achieved by reducing of the PHEMA molecular weight to < 2,500 Da7. In this respect, controlled copolymerizations, such as reversible addition-fragmentation chain transfer (RAFT)8,9 or atom transfer radical polymerization (ATRP)10 were natural choices for the preparation of degradable PHEMA. An additional advantage of copolymerization is the easy introduction of functional groups, which are subsequently available for further attachment of biomolecules. Degradable polymers were also prepared by radical polymerization in the presence of crosslinkers that contain degradable bonds, such as (meth)acryloylated dextran11, polyglutamic acid12, caprolactone13, anhydrides14, or monomers containing disulfide bonds15, e.g., N,N′bis(methacryloyl)-L-cystine16,

bis(2-methacryloyl)oxyethyl

disulfide17,

and

N,N′-

bis(acryloyl)cystamine18. Alternatively, hydrogels were formed by the crosslinking of reactive polymers, e.g., poly(ethylene glycol)19, dextran20, or chitosan21, with degradable bonds, such as disulfide bridges, peptide linkers22, hydrazone, and esters. The reductive degradability was preferred because hydrolytically degradable hydrogels are difficult to store, and enzymatic degradation requires expensive peptide sequences. A concept, which is not commonly used for the preparation of reductively degradable disulfide scaffolds, involves the controlled radical polymerization of reactive methacrylates or methacrylamides in solution, which is followed by gelation of the copolymers by polymer-analogous reactions. Introducing thiols into polymers can be accomplished by polymerization of protected thiols or cyclic precursor e. g. N-(acryloyl) homocysteine-γ-thiolactone which is not releasing side-products upon deprotection reactions.23

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The novelty of this work consist in preparation of well-defined reactive water soluble lowmolecular-weight poly(2-hydroxyethyl methacrylate-co-(2-acetylthio)ethyl methacrylate-co-2methacryloyloxyethyl phosphorylcholine) [P(HEMA-ATEMA-MPC)] by reversible additionfragmentation chain transfer (RAFT) polymerization, followed by the controlled deprotection of SH groups which allowed to structure hydrogel during controlled gelation in the presence of a porogen, Above that, 2-methacryloyloxyethyl phosphorylcholine (MPC) was used as a comonomer to facilitate the solubility of PHEMA. MPC also acts as a biomimetic component of Omafilcon A artificial cornea24, intraocular lenses25, cardiovascular stents26, implantable blood pumps27, and artificial hip joints28,29. Because living tissues, such as nerves, muscles, tendons, ligaments, and dentin, have an oriented tubular or fibrous structure, the organization and regeneration of these tissues in hydrogel scaffolds with an oriented porosity can be beneficial. To introduce an oriented porosity in the hydrogel, needle-like sodium acetate crystals were used as a porogen because (NH4)2SO430 and ammonium oxalate31 were disqualified; due to their large size and reductive properties in this study. Moreover, the in vitro biocompatibility and the cytotoxicity of the polymers were investigated using human mesenchymal stem cells (MSCs).

EXPERIMENTAL

Chemicals 2-Hydroxyethyl methacrylate (HEMA; Wichterle&Vacík; Prague, Czech Republic) was vacuum-distilled

before

use.

The

inhibitor

was

removed

from

zwitterionic

2-

methacryloyloxyethyl phosphorylcholine (MPC; TCI; Portland, OR, USA) on an aluminum 4 ACS Paragon Plus Environment

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oxide column. 4,4′-Azobis(4-cyanovaleric acid) (ACVA), 2,2′-azobis(2-methylpropionitrile) (AIBN), 4-cyano-4-(phenylcarbonothioylthio)pentanoic acid (CPTPA; chain transfer agent), 2bromoethanol,

triethylamine,

N,N-ethyldiisopropylamine,

potassium

thioacetate,

2,2′-

dithiodipyridine (DTP), methacryloyl chloride, L-cysteine, and 1-phenyl-3-pyrazolidinone (Sigma-Aldrich; St. Louis, MO, USA) were used as received. Ethanol, methanol, acetone, dichloromethane, toluene, diethyl ether, and anhydrous sodium acetate (recrystallized in water) were obtained from LachNer (Neratovice, Czech Republic). Cellulose acetate butyrate (CAB; acetyl/butyryl groups 35/15, Mn = 100,000) was obtained from Eastman Chemical Company (Kingsport, TN, USA). For cell cultivation, alpha-minimum essential medium (MEM; EastPort; Prague, Czech Republic), platelet lysate (IKEM; Prague, Czech Republic), trypsin/ethylenediaminetetraacetic acid (EDTA) solution (Life Technologies; Prague, Czech Republic), gentamicin (Sandoz; Prague, Czech Republic), antibiotic antimycotic solution (Sigma), and L-glutamine (Sigma) were used. AlamarBlue assay was purchased from ThermoFisher (Waltham; MA, USA). Flow cytometry (FACSAria™; Becton Dickinson; San Jose, CA, USA) was used for the detection of the following antibodies: CD34, CD45, and CD105 (Exbio; Vestec, Czech Republic), CD29, CD73, CD90, CD31, HLA-ABC, and HLA-DR (BD Pharmingen; San Jose, CA, USA), and CD133 (Miltenyi Biotec; Bergisch Gladbach, Germany). Triton X-100, bovine serum albumin (BSA), and immunofluorescent stains Chemiblocker and Alexa-Fluor 568 phalloidin were purchased from Merck Millipore (Prague, Czech Republic).

Synthesis of 2-bromoethyl methacrylate



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A

mixture

of

2-bromoethanol

(15.5

g),

N,N-ethyldiisopropylamine

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(13

g),

and

dichloromethane (100 ml) was cooled in a bath of dry ice and ethanol at -78 °C, and methacryloyl chloride (10.5 g) was added. The solution was slowly heated to RT and kept at this temperature for 3 h. Dichloromethane was evaporated, 1-phenyl-3-pyrazolidinone inhibitor (100 mg) was added, and the resulting viscous substance was vacuum-distilled. A fraction, b. p. 49 °C (1066 Pa), was collected (11.8 g, 61 %). 1H NMR in CDCl3: δ (ppm) 1.95 (s, 3H), 3.7 (m, 2H), 4.4 (m, 2H), 5.6 (dd. H), and 6.1 (dd. H).

Synthesis of (2-acetylthio)ethyl methacrylate A previously described procedure was slightly modified32. Briefly, 2-bromoethyl methacrylate (10 g) and potassium thioacetate (6 g) were dispersed in acetone, 1-phenyl-3-pyrazolidinone inhibitor (100 mg) was added, and the reaction mixture was stirred at RT for 24 h. The resulting suspension was filtered, and the acetone evaporated; the residuum was redispersed in dichloromethane, washed in water, and dried by MgSO4. Dichloromethane was removed on a rotary evaporator, and the crude product was vacuum-distilled. A fraction, b. p. 90 °C (400-533 Pa), was collected (4.9 g, 50 %). 1H NMR in CDCl3: δ (ppm) 1.95 (s, 3H), 2.35 (s, 3H), 3.25 (t, 2H), 4.2 (t, 2H), 5.6 (dd., H), and 6.1 (dd., H).

Preparation of low-molecular-weight PHEMA, P(HEMA-MPC), P(HEMA-ATEMAMPC), and P(HEMA-ATEMA-MPC)-SH Low-molecular-weight PHEMA, P(HEMA-MPC), and P(HEMA-ATEMA-MPC) were prepared by reversible addition-fragmentation chain transfer (RAFT) polymerization. In a typical 6 ACS Paragon Plus Environment

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experiment, a solution of HEMA (2.738 g), ATEMA (0.365 g), MPC (0.574 g), CPTPA (0.04 g), and ACVA (0.02 g) in ethanol (8.5 ml) was purged with argon for 30 min and polymerized at 70 °C for 12 h. PHEMA and P(HEMA-MPC) were prepared analogously. Advantages of ethanol as polymerization solvent are high purity, favorable chain transfer constant and boiling point, and ability to solubilize the product. The resulting polymer was precipitated in diethyl ether and redispersed in methanol; AIBN (1 g) was added, and the mixture was heated at 60 °C for 2 h to remove dithiobenzoic groups. The mixture was cooled at -20 °C, and the precipitate was collected and dissolved in methanol. The procedure was repeated until a white product was obtained. Finally, the P(HEMA-ATEMA-MPC) was precipitated in diethyl ether and vacuumdried. The polymer with the deprotected thiol groups (P(HEMA-ATEMA-MPC)-SH) was prepared by stirring P(HEMA-ATEMA-MPC) in an aqueous solution (100 ml) of triethylamine (0.35 ml) at RT for 10 h under argon atmosphere; the polymer was dialyzed under the same atmosphere and lyophilized.

Preparation of nonporous hydrogels (NPH) Nonporous hydrogels were prepared from P(HEMA-ATEMA-MPC) containing various amounts of ATEMA. The polymers (100 mg) containing protected thiol groups and stoichiometric amounts of DTP were dissolved in a 2-ml Eppendorf centrifugation tube in a mixture of methanol (0.08 ml) and water (0.02 ml). When complete dissolution was achieved, triethylamine (0.035 ml) was added to hydrolyze the protecting group and a mixture was shorty vigorously stirred. The reaction continued at RT for 2 days. The resulting NPH hydrogels were washed with ethanol to remove DTP and water.



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Preparation of hydrogels with oriented porosity (PH) Polyethylene syringes (5 ml) equipped with a stainless mesh (40 µm) were filled with hot (75 °C) filtered sodium acetate solution (7 mol/l) and slowly immersed in the ethanol/dry ice bath (-75 °C). The syringes were warmed up to 10 °C, and excessive sodium acetate solution was removed under vacuum; the crystals were quickly washed with acetone and dried under a flow of air. To increase the porosity, one syringe was repeatedly filled with sodium acetate solution. Alternatively, sodium acetate trihydrate crystals were sedimented in a halve-cut tube (1.24 mm in diameter) and transferred into the syringe. To protect the crystals from dissolution, CH3COONa (0.6 g) was added to the polymer solution, or the crystals were impregnated by 5 wt.% CAB in acetone. The syringe was filled with a honey-like mixture consisting of P(HEMAATEMA-MPC) Run 8 (2 g), methanol (1.6 ml), water (0.4 ml), DTP (90 mg), and triethylamine (0.7 ml). The syringe was closed by a pressure cap, air bubbles between the crystals were removed by centrifugation, and the mixture was reacted at RT for 2 days. The resulting porous hydrogel was denoted as PH-S (sedimentation method). The hydrogels obtained by filling the syringes with sodium acetate solution and freezing at -75 °C were denoted as PH-F1 and PH-F2 for single and double filling, respectively. The hydrogels were removed from the syringes and cut to 1-mm thick discs, which were washed with acetone to remove CAB from pores, ethanol, phosphate buffered saline (PBS) to reduce swelling of hydrogels during solvent exchange, and distilled water.

In vitro degradation of the hydrogels Degradation of NPH-1 - 6, PH-S, PH-F1, and PH-F2 was investigated according to a previously described procedure15. The hydrogels (100 mg of dry weight) were immersed in a 8 ACS Paragon Plus Environment

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solution of L-cysteine (3.3 mmol/l) in PBS (50 ml) at 37 °C and weighed at periodic intervals. The degree of degradation (DD) was calculated according to Eq. 1, where m and m0 are the weights of the swollen hydrogel at a specific time and at the beginning of the experiment, respectively. The porous hydrogels were degraded in different concentrations of L-cysteine (0.33 and 3.3 mmol/l), which was accompanied with the daily replacement of the old L-cysteine solution for a fresh L-cysteine solution.

DD = (m-m0)/m0

(1)

Characterization The P(HEMA-ATEMA-MPC) hydrogel discs were analyzed by a low-vacuum scanning electron microscope (SEM) with a Quanta 200 FEG microscope (FEI; Brno, Czech Republic) at -10 °C and 100 Pa. The composition of the hydrogels was determined with a Perkin-Elmer 2400 CHN elemental analyzer (Waltham, MA, USA). The sulfur content was determined by titration with barium perchlorate. The infrared spectra were recorded by a Nexus Nicolet 870 FTIR spectrometer (Ramsey, MN, USA) equipped with a liquid nitrogen-cooled mercury cadmium telluride detector using a Golden Gate single reflection ATR cell (Specac; Slough, UK). The cumulative pore volume, the porosity, and the average pore size of the dry hydrogels were determined using Pascal 140 and 440 mercury porosimeters (Thermo Finigan; Rodano, Italy) at pressure intervals of 0–400 kPa and 1–400 MPa (pore diameter 0.004-116 µm). The specific surface area was determined by dynamic desorption of nitrogen at -196 °C using a Gemini VII 2390 Analyzer (Micromeritics; Norcross, GA, USA). The composition of the monomers and kinetics of the hydrogel formation in D2O/methanol-d4 mixture were determined using a 1H 9 ACS Paragon Plus Environment


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NMR Bruker DPX 300 and an Avance III 600 spectrometer (Billerica, MA, USA), respectively. Chemical shifts were calibrated to the HDO signal at δ = 4.80 ppm. The kinetics was measured from the DTP peaks in the aromatic region of the spectra polymerization mixture at 0.5 and 65 h using signals 1 and b (Figure 1).

Figure 1. 1H NMR spectra of 2,2'-dithiodipyiridine and pyridine-2-thione during deprotection and oxidation of P(HEMA-ATEMA-MPC) Run 8 for (A) 0.5 and (B) 65 h. The porosity (P) of the hydrogels was determined from the volume (V), weight (w), and density (ρ = 1.45 g/ml) of sodium acetate trihydrate, according to Eq. 2.

ܲൌ

౭ ಙ

ቀ୚ି ቁൈଵ଴଴ ୚

(%)

(2) 10

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The hydrodynamic particle size after hydrogel degradation was measured using dynamic light scattering on a Zetasizer Nano Instrument ZEN 3600 (Malvern Instruments; Malvern, UK). The molecular weight and polydispersity of the polymers were determined using a Shimadzu HPLC size exclusion chromatograph (Tokyo, Japan) equipped with a UV−VIS diode array, a refractive index OptilabrEX, and multiangle light scattering DAWN EOS detectors (Wyatt; Santa Barbara, CA, USA). A TSK SuperAW3000 column and methanol/sodium acetate buffer (80/20 v/v) eluent (pH 6.5) at a flow rate of 0.6 ml/min were used. Solvent regains SR (g/g) were calculated according to Eq. 3, where wd and ww are the weights of the dry and swollen hydrogel, respectively.

SR = (ww - wd)/wd

(3)

Chemorheological determination of the gel point Rheological measurements were carried out using a Bohlin Gemini HR Nano oscillatory rheometer (Malvern, UK) equipped with coaxial cylinders C25 integrated with a Peltier measuring table for temperature control. The viscoelastic response of the sample due to gelation was monitored based on changes in the storage (G’) and loss (G’’) shear moduli and the loss factor (tan δ) during the crosslinking reaction. The deformation responses of the in situ crosslinking were collected within a small strain region at six frequencies between 0.4 and 3.2 Hz using the multiple-frequency mode. First, the initial curing region up to the liquid-solid transition was monitored by the stress-control mode, and then, when the system passed through 11 ACS Paragon Plus Environment


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the gel, the mode of deformation control was switched to the strain-control mode. This ensured that the condition of the so-called linear viscoelastic region applied throughout the entire curing cycle, and at the same time, the applied and measured forces and strains were relevant. For selecting the measuring strain, which was 10-3, the linear viscoelastic region was determined, and the strain value was used. The frequency-dependent dynamic shear moduli G’ and G’’ and the loss factor (tan δ = G’’/G’) were plotted as functions of the gelation time. The gel time was determined as the time at which G’ and G’’ in the log-log frequency plot showed a congruent behavior, according to a previously established theory33. This time also coincided with the range at which the tan δ did not depend on the measuring frequency. To prepare the hydrogel, the reaction components were first mixed in a vial, and then the exact volume of the liquid (13 ml) was transferred into the cylindrical cup using a syringe fitted with a needle. The reaction time started when the components were mixed, and the reaction proceeded at 18 °C.

Cell culture experiments Human mesenchymal stem cells (MSCs) were isolated from the human umbilical cord tissue. Fresh umbilical cords were collected from healthy full-term neonates after spontaneous delivery with the informed consent of the donors using the guidelines approved by the Institutional Ethics Committee at University Hospitals in Pilsen and Prague, Czech Republic. Approximately 10-15 cm per umbilical cord was aseptically transported into sterile PBS with antibiotic–antimycotic solution at 4 °C. After removal of blood vessels, the remaining tissue was chopped into small pieces (1-2 mm3), transferred to culture dishes containing the complete alpha-MEM medium supplemented with 5 % platelet lysate and gentamicin (10 µg/ml), and cultivated at 37 °C in a humidified 5 % CO2 atmosphere. The explants were left undisturbed for up to 5 days to allow the 12 ACS Paragon Plus Environment

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migration of cells, at which point part of the medium was replaced. On day 10, the explants were removed from the culture dishes, and the remaining adherent cells were cultured for 3 weeks or until 90 % confluence was achieved. The cells were passaged after digestion with 0.25 % trypsin/EDTA solution and re-seeded at a concentration of 5 × 103/cm2 on 80 cm2 culture flasks. The medium was changed two times a week. Cells of the 3rd passage were identified by flow cytometry to confirm their purity using antibodies against human antigens CD34-, CD45-, CD31-, HLA-DR-, CD105+, CD29+, CD73+, CD90+, CD133+, and HLA-ABC+. Data were analyzed using the BD FASCDiVa software (FACSAria™). To evaluate MSC proliferation, sterile hydrogel discs were cut into quarters and placed into a 48-well plate with the MSC culture medium. A suspension of 30,000 MSCs was seeded per disc and incubated at 37 °C in a humidified 5 % CO2 atmosphere. The viability of the MSCs grown on the tested hydrogels was determined after 1, 3, and 7 days of the culture using an AlamarBlue® assay. AlamarBlue® reagent was added to a culture medium and incubated at 37 °C for 3 h. The medium was then transferred to a 96-well plate, and the fluorescence was measured on a Tecan-Spectra spectrophotometer reader (Mannedorf, Switzerland) using the 560ex nm/590em nm filter setting. After the measurement, the fresh culture medium was added to the cells, and the cells on the hydrogels were further incubated. Each type of the hydrogel was seeded in quadruplets. Lyophilized P(HEMA-ATEMA-MPC)-SH and P(HEMA-ATEMA-MPC) Run 8 were dissolved in the cell culture medium filtered using a 0.22 µm syringe filter (TPP; Trasadingen, Switzerland) at concentrations of 0, 0.25, 0.5, 1, 2.5, and 5 wt.% and added to a MSC culture seeded in a 96-well plate (3000 cells per well) for 1 day. Cell proliferation was determined using the AlamarBlue® reagent, as described above.



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For fluorescent staining, MSCs grown on the hydrogel discs were fixed in 4 % paraformaldehyde in 0.1 M PBS for 15 min, washed with PBS, treated with 0.5 % Triton X-100 in PBS, and incubated with Alexa-Fluor 568 phalloidin (1:300). The discs were observed using an Axioscop 2 light microscope (Zeiss; Jena, Germany).

RESULTS AND DISCUSION

Preparation of P(HEMA-ATEMA-MPC) P(HEMA-ATEMA-MPC) was designed to be soluble under physiological conditions, i.e., in PBS, water, or alpha-MEM. The molecular weight of the polymer must be < 45 kDa, which is the molecular-weight threshold limiting glomerular filtration34. To fulfill this requirement, HEMA was copolymerized with the highly hydrophilic zwitterionic monomer 2methacryloyloxyethyl phosphorylcholine (MPC). To introduce reactive thiol groups in the polymer, (2-acetylthio)ethyl methacrylate (ATEMA) was synthetized and used as a second comonomer. Reactive copolymers with a narrow molecular weight distribution were obtained by RAFT copolymerization of HEMA, ATEMA, and MPC in ethanol using ACVA as the initiator and CPTPA as the chain transfer agent (Figure 2).


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Figure 2. RAFT copolymerization of 2-hydroxyethyl methacrylate (HEMA), 2-(acethylthio)ethyl methacrylate (ATEMA) and 2-methacryloyloxyethyl phosphorylcholine (MPC) and removal of terminal dithiobenzoic group. R1 -OH, R2 -SCOCH3 and R3 -OPO3-C2H4N+(CH3)3.

The effect of the ATEMA amount on the properties of P(HEMA-ATMA-MPC) was determined to prepare polymers with various contents of reactive thiol groups. P(HEMAATEMA-MPC) served as a precursor for the formation of hydrogels with various degrees of crosslinking. Increasing the concentration of ATEMA increased the molecular weight and slightly broadened the polydispersity of P(HEMA-ATEMA-MPC) (Table 1).

Table 1. Effect of amount of ATEMA on molecular weight and composition of P(HEMAATEMA-MPC). Composition of monomers C (wt.%) H (wt.%) N (wt.%) in polymerization feed Mw Run PD HEMA MPC ATEMA (kDa) Dtm. Calc. Dtm. Calc. Dtm. Calc. (wt.%) (wt.%) (wt.%) 1 83.1 15.5 1.4 42 1.08 52.8 53.6 7.8 7.6 0.9 0.7 2 81.7 15.5 2.8 43 1.08 52.0 53.5 7.8 7.6 0.7 0.7 3 79.0 15.5 5.4 44 1.10 52.7 53.4 7.8 7.6 0.1 0.7 4 76.3 15.5 8.2 46 1.10 51.8 53.4 7.7 7.5 0.9 0.7 5 73.6 15.5 10.9 47 1.10 52.4 53.2 7.6 7.5 0.9 0.7

S (wt.%) Dtm. Calc. 0.6 0.5 0.9 1.2 1.6

0.2 0.5 0.9 1.3 1.9

Monomers/CPTPA/ACVA = 183.3/2/1 w/w/w; HEMA – 2-hydroxyethyl methacrylate; MPC – 2-methacryloyloxyethyl phosphorylcholine; ATEMA – 2-(acethylthio)ethyl methacrylate; CPTPA – 4-cyano-4-(phenylcarbonothionylthio)pentanoic acid; ACVA - 4,4′-azobis(4cyanovaleric acid); PD – polydispersity; Dtm. – determined; Calc. – calculated.

Because the influence of the thioacetate group on the molecular weight of the polymers should be negligible32,35,36, the above-mentioned effect could be ascribed to a presence of impurities in



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ATEMA. After three months of storage, P(HEMA-ATEMA-MPC) became insoluble. The problem was solved by heating the polymer with AIBN in ethanol, which removed the dithiobenzoic groups. Water-soluble P(HEMA-ATEMA-MPC) with a sufficient amount of reactive groups available for future gelation preferably contained ~15 wt.% of MPC and more than 2.8 wt.% of ATEMA. The molecular weight and polydispersity of the P(HEMA-ATEMAMPC) were controlled by changing the monomers/ACVA ratio with constant CPTPA/ACVA and HEMA/ATEMA/MPC ratios of 2/1 (w/w) and 7.4/1.54/1 (w/w/w), respectively (Table 2). Results from the elemental analysis of P(HEMA-ATMA-MPC) were compared with the theoretical contents of C, H, N, and S to determine the composition of the copolymers (Table 1, 2). The contents of the analyzed elements roughly corresponded to the theoretical amounts.

Table 2. Effect of amount of CPTPA and ACVA on molecular weight and composition of P(HEMA-ATEMA-MPC).

CPTPA ACVA Mw Run (g) (g) (kDa) 6 7 8 9 10

0.01 0.02 0.04 0.06 0.08

0.005 0.01 0.02 0.03 0.04

259. 131 49 35 28

C (wt.%) PD 1.10 1.12 1.08 1.09 1.09

H (wt.%)

N (wt.%)

S (wt.%)

Dtm.

Calc.

Dtm.

Calc.

Dtm.

Calc.

Dtm.

Calc.

52.8 53.1 52.6 53.1 54.2

53.2 53.2 53.2 53.2 53.2

8.1 8.4 8.2 8.3 8.6

7.5 7.5 7.5 7.5 7.5

0.7 1.0 0.9 0.9 0.9

0.7 0.7 0.7 0.7 0.7

1.8 1.7 1.6 1.7 1.7

1.7 1.7 1.7 1.7 1.7

Composition of polymerization mixture: 2.74 g HEMA, 0.57 g MPC, 0.37 g ATEMA; HEMA – 2-hydroxyethyl methacrylate; MPC – 2-methacryloyloxyethyl phosphorylcholine; ATEMA – 2-(acethylthio)ethyl methacrylate; CPTPA – 4-cyano-4-(phenylcarbonothionylthio)pentanoic acid; ACVA - 4,4′-azobis(4-cyanovaleric acid); PD – polydispersity; Dtm. – determined; Calc. – calculated.


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The chemical compositions of PHEMA, P(HEMA-MPC), and P(HEMA-ATEMA-MPC) were investigated by FTIR spectroscopy (Figure 3). The peak at 3430 cm-1 in the spectrum of PHEMA was ascribed to the O-H stretching vibration; peaks at 2886, 2946, and 2986 cm-1 belonged to the antisymmetric and symmetric stretching vibrations of CH3, CH2, and CH, respectively. The peak at 1718 cm-1 was attributed to the C=O stretching vibration, the peak at 1452 cm-1 was attributed to the in-plane bending of CH2, the peak at 1390 cm-1 corresponded to the CH2 twist and rock, and the peaks at 1076 and 750 cm-1 were attributed to the C-O-C stretching vibration and the -CO- out-of-plane bending, respectively37. New peaks at 790 and 960 cm-1 appeared in the spectrum of P(HEMA-MPC), which were attributed to antisymmetric stretching of P(OC)2 and symmetric stretching of N+(CH3)3, respectively38,39. The FTIR spectrum of (PHEMA-ATEMAMPC) was characterized by peak at 624 cm-1, which is ascribed to the C(O)-S vibration40,41 of ATEMA (Figure 3).

Figure 3. FTIR spectra of (1) PHEMA, (2) P(HEMA-MPC), (3) P(HEMA-ATEMA-MPC), and (4) PH-F1 hydrogel.



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Preparation of reductively degradable hydrogels Degradable PHEMA hydrogels were prepared by deprotection of thiol groups of P(HEMAATEMA-MPC) followed by oxidation with 2,2'-dithiopyridine (Figure 4); this reaction is often used for coupling of proteins42. The reactions should be slow to provide sufficient time for the manipulation of the solidifying mixture, to enable the filling of the syringe with a viscous polymer solution and to remove air bubbles. Cleavage of acetyl groups is usually performed with NaOH, but thiol groups can also be deprotected with other bases or acids43. In this work, triethylamine was selected because it enables a slow deprotection of thiols and is well-miscible with aqueous and organic solvents.

Figure 4. Scheme of hydrogel formation by the simultaneous basic deprotection of thiol groups of P(HEMA-ATEMA-MCP) and oxidation with 2,2'-dithiopyridine (DTP).

To introduce an oriented porosity in the hydrogels, sodium acetate trihydrate was used as a porogen, which leaves pores after its dissolution when hydrogel formation is completed. Sodium acetate trihydrate has a needle-like shape, and it does not reduce disulfide bridges, which is in contrast to the ammonium oxalate used in our previous reports31,44. Moreover, thin sodium 18 ACS Paragon Plus Environment

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acetate trihydrate needles can grow from a single nucleation center, which is advantageous for the introduction of an oriented porosity. Slight disadvantages include the high solubility of the sodium acetate in a water/ethanol mixture and its low melting point (60 °C). Three degradable hydrogels with an oriented porosity were prepared. The first hydrogel, denoted as PH-S, was prepared by the sedimentation of sodium acetate trihydrate in a half-cut tube, according to a previously published procedure31. To protect the crystals from dissolution in methanol, they were impregnated with a solution of CAB in acetone and dried. Because the introduction of an oriented porosity in the hydrogel through this method was laborious and time consuming, a new procedure was developed for the preparation of porous hydrogels. The process was inspired by the preparation of porous hydrogels using a unidirectional freezing technique45. An organized structure was achieved by crystal growth in the direction of the temperature gradient. Thus, the second and third hydrogels were obtained with different numbers of fillings with the crystals; the hydrogels were denoted as PH-F1 and PH-F2, respectively. The porosity of PH-S (~53 vol.%) calculated from the volume, weight, and density of the sodium acetate trihydrate crystals differed from that obtained from mercury porosimetry (79.6 vol.%; Table 3). The oriented pore structure of PH-S was confirmed by SEM (Figure 5 A, B).



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Figure 5. SEM micrographs of (A, C, E) perpendicular and (B, D, F) parallel cross-sections of (A, B) PH-S, (C, D) PH-F1, and (E, F) PH-F2 hydrogels. Scale bar 500 µm.

The pore size obtained from SEM (51-200 µm) roughly corresponded with that from mercury porosimetry (75 µm). The porosities of PH-F1 and PH-F2, according to mercury porosimetry (70.0 and 70.1 vol.%), also did not agree with the values calculated from the amount of crystals in the reaction feed (50 and 75 vol.%). The discrepancy between the results from mercury porosimetry and other methods can be attributed to limitations of the former method, which measures pores only in range 4-100 µm, and to effects, such as the formation of cracks and shattering of samples during measurements. Thick polymer walls were clearly visible in the SEM micrographs of a cross-section of PH-F1 (Figure 5 C, D) compared to PH-F2 (Figure 5 E, F). 20 ACS Paragon Plus Environment

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The specific surface areas of both hydrogels were low (0.35-0.40 m2/g) due to the large pore sizes. The diameters of the PH-F2 pores were 45-75 and 65 µm, according to SEM and Hg porosimetry, respectively. In contrast, the diameters of the PH-F1 pores were 200-1,000 and 59 µm, according to SEM and Hg porosimetry, respectively. Such a pore size is suitable for future cell accommodation. The variability of the pore sizes and shapes is caused by the crystal origins. Because crystallization of sodium acetate for PH-S was induced by cooling, and the crystals were left to sediment, the pores in the hydrogel were regular, and they exhibited a slightly rounded shape due to surface erosion. The size of the crystals for the PH-F1 and PH-F2 hydrogels was influenced by the temperature gradient and crystallization velocity. Crystals for PH-F2 were smaller and more regular than those for PH-F1, which is probably due to the partial dissolution of primary crystals during the second loading run of the hot sodium acetate solution. Table 3. Porous properties of hydrogels. Hydrogel PH-S PH-F1 PH-F2

DSEM (µm) 51-200 200-1000 45-75

DHg (µm) 75 59 65

P (vol.%) 79.6 70.1 70.0

SBET (m2/g) 0.35 0.02 0.40

DSEM and DHg – pore diameter according to SEM and Hg porosimetry, respectively; P porosity; SBET – specific surface area according to dynamic desorption of nitrogen.

The FTIR spectrum of PH-F1 hydrogel confirmed the successful deprotection of thiol groups of (PHEMA-ATEMA-MPC) with triethylamine due to the disappearance of the peak at 624 cm-1. (Figure 3). Differently crosslinked nonporous and porous hydrogels were prepared, and their water, ethanol, and cyclohexane regains were measured (Table 4). The water regain was always higher than the ethanol regain, which was due to the presence of hydrophilic MPC units in the hydrogel. 21 ACS Paragon Plus Environment


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Table 4. Solvent regain of nonporous (NPH) and porous PHEMA hydrogels (PH).

Hydrogel

P(HEMA-ATEMAMPC)

DTP/polymer (g/g)

Water regain (g/g)

EtOH regain (g/g)

NPH-1 NPH-2 NPH-3 NPH-4 NPH-5 NPH-6 PH-F1 PH-F2 PH-S

Run 2 Run 2 Run 3 Run 4 Run 8 Run 5 Run 8 Run 8 Run 8

0.015 0.016 0.030 0.038 0.045 0.060 0.045 0.045 0.045

4.51 3.37 2.84 3.19 2.19 1.99 8.00 11.10 7.30

0.62 0.40 0.59 0.57 0.47 0.59 2.50 3.60 2.50

Cyclohexane regain (g/g) 0.33 0.22 0.11 0.11 0.11 0.11 1.70 2.20 1.80

HEMA – 2-hydroxyethyl methacrylate; ATEMA – 2-(acethylthio)ethyl methacrylate; MPC – 2-methacryloyloxyethyl phosphorylcholine; DTP – 2,2'-dithiopyridine.

Both the water and cyclohexane regains in the hydrogels roughly corresponded to the amount of ATEMA. However, the ethanol regain was independent of the ATEMA content in the hydrogels; this is probably due to the inability to remove all the water from the hydrogels to achieve equilibrium swelling. The kinetics of the hydrogel formation, namely, the decreasing concentration of DTP in the reaction mixture, was measured with 1H NMR spectroscopy (Figure 6). Coupling of the thiol groups of the neighboring polymer chains via DTP resulted in the formation of disulfide bridges. Since each thiol group generates one pyridine-2-thione molecule, the quantification was straightforward. The DTP was completely consumed after 5-h of the reaction.


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Figure 6. Conversion of 2,2'-dithiodipyiridine to pyridine-2-thione during deprotection and cross-linking of P(HEMA-ATEMA-MPC) Run 8, according to 1H NMR spectroscopy.

Chemorheological measurement of degradable hydrogel formation To investigate the cross-linking of P(HEMA-ATEMA-MPC) Run 8 in terms of its mechanical properties, the rheological behavior during the reaction was investigated. The storage (G’) and loss (G’’) shear moduli changed with the reaction time. At the beginning of the reaction, when the mixture formed a low-viscous liquid, the loss modulus was higher than the storage modulus. At a specific reaction time, the storage modulus became larger than the loss modulus, which was an indication of the liquid-solid transition that is attributed to gelation (Figure 7).



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Figure 7. Changes of storage (G’) and loss (G’’) shear moduli and loss factor (tan δ) during the cross-linking of P(HEMA-ATEMA-MPC) Run 8 as a function of time at different frequencies: 0.4, 0.8, 1.2, 1.6, 2.0, and 3.2 Hz.

Above the gel point, the mixture exhibited a viscoelastic response, which is typical of solid materials and corresponds to a covalently crosslinked polymer at its rubbery state with the prevailing storage modulus. Both moduli continued increasing beyond the gel time as the network structure was developed, and the number of network chains increased further. The G’G’’ crossover is the empirical criterion of the gel point33,46. Because the crossover gelation criterion is known to somewhat depend on the deformation frequency, gelation was measured at a frequency range of 0.4-3.2 Hz. The positions of all the crossovers indicated that the gel time occurred between approximately 63 and 86 min (Figure 7). The generally accepted criterion for determining the gel time independent of the frequency is the plot of tan δ vs. the reaction time for a set of frequencies. According to this parameter, the gel point was determined as 72 min. Next, parallel courses of both moduli with the frequency in a log-log plot were investigated. The 24 ACS Paragon Plus Environment

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closely parallel courses of G’(f) and G’’(f) had slopes of ca. 0.76 at the reaction time of 71 min. This time corresponded with the one determined according to the Winter-Chambon criterion of tan δ independency of frequency33. The fully crosslinked nonporous hydrogel, NPH-5 (c.f., Table 4) of equilibrium water regain SR = 2.2 g/g had a shear modulus of approx. 17 kPa (independent of frequency in the range 10-3-102 Hz). This value was close to the shear modulus values of covalently crosslinked PHEMA hydrogels of the matching water regain47 (SR = 2-2.3 g/g).

Reductive degradation of hydrogels Two concentrations of L-cysteine (0.33 and 3.3 mmol/l in PBS) were used for degradation of the hydrogels. The first value simulated the concentration of L-cysteine in Dulbecco’s Modified Eagle’s Medium and the total thiol levels in human plasma containing L-cysteine (200 µM), Lcysteine-L-glycine (100 µM), homocysteine (7 µM), γ-L-glutamyl-L-cysteine (5 µM), and glutathione (4 µM). Concentrations of low-molecular sulfides in their reduced form in the plasma are, however, significantly lower (L-cysteine 9 µmol/l, glutathione 1.5 µmol/l, L-cysteinyl glycine, 1.5 µmol/l, L-homocysteine 0.15 µmol/l)15,48. The second concentration (3.3 mmol of Lcysteine/l) was used to accelerate the experiments. It can be assumed that in vivo hydrogel degradation will be slower than that at 0.33 mmol/l, although the degradation can be accelerated due to the SH/SS exchange reaction, where thiol only acts as a catalyst. Table 5 and Figure 8 show the degree of degradation and time required for dissolution of the hydrogels.



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Table 5. Degradation of PH and NPH hydrogels.

Hydrogel NPH-5 PH-F1 PH-F2 PH-S

Time 3.3 mmol of L-cysteine/l 0.33 mmol of L-cysteine/l 31 h 86 d 24 h 86 d 9h 57 d 27 h 86 d

Figure 8. Degradation of NPH-1 - 6 hydrogels in 3.3 mmol of L-cysteine/l PBS.

Negative degrees of degradation (DD) correspond to a decrease of the hydrogel weight, and DDs were caused by increased water regain due to a decreasing number of disulphidic bridges as a result of degradation. The low-crosslinked NPH-1 had poor mechanical properties, and therefore, it rapidly decomposed in 3.3 mmol of L-cysteine/l during 30 min. The degradation of the nonporous gels NH-1-8 was prolonged from 0.5 to 53 h as the crosslinking of the hydrogel increased (Figure 8). The degradation of the porous hydrogels was difficult to measure due to the high amount of water within the pores because only ca. 10 wt.% of the swollen hydrogel consisted of the polymer. As the degree of crosslinking decreased, the hydrogel exhibited a 26 ACS Paragon Plus Environment

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tendency to swell, increase its pore size, and lose its shape. The time for complete dissolution is recorded in Table 5. Light micrographs of the PH-S and PH-F1 degradation products in Lcysteine solution were taken over time, and the mass loss of both hydrogels was clearly visible (Figure 9 A, B). Finally, the dynamic light scattering of the degradation products was measured (Figure 9 C). Notably, the PH-S degraded to almost uniform particles.

Figure 9. Light micrographs of the degradation products of (A) PH-S and (B) PH-F1 in 0.33 mmol of L-cystein/l (rigt) compared to those in PBS (left); (C) DLS of residual solution after complete degradation of (1) PH-F1 and (2) PH-S.

In contrast, PH-F1 disintegrated into particles with broad particle size distribution. This can be caused by the presence of residual CAB in the hydrogel. PH-S, PH-F1, and NH-8 fully degraded in 0.33 mmol of L-cysteine/l within 85 days. PH-F2 degraded significantly more rapidly, which is probably due to its highly-organized pore structure (Table 5). Cell experiments were



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conducted with PH-F1 and PH-F2 hydrogels to avoid possible inconsistencies caused by CAB in PH-S.

Cell experiments The cell adhesive properties of both PH-F1 and PH-F2 hydrogels were investigated in MSC culture for 1, 3, and 7 days. The cells attached to the PH-F1 hydrogel surface, but cell spreading was restricted due to the lack of active adhesive sites (Figure 10 A, B). The cells displayed a round shape instead of a flattened morphology, which is common, for instance, on the tissue culture plastic or on hydrogels modified with cell-adhesive peptides31,42,49,50. Despite the limited adhesion, the cells remained attached, and they proliferated in the form of clusters on both the PH-F1 and PH-F2 hydrogels after 7 days of the culture, which confirmed the biocompatibility and non-cytotoxicity of the material (Figure 10 C).


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Figure 10. Fluorescence micrographs of phalloidin-stained MSCs seeded on PH-F1 for (A) 1 and (B) 7 days. (C) MSCs proliferation on PH-F1 and PH-F2 hydrogels (dark and light columns, respectively) measured by the AlamarBlue assay after 1, 3, and 7 days of culture (mean ± standard deviation, n = 4). Scale bar 100 µm.

The toxicity of P(HEMA-ATEMA-MPC) Run 8 and P(HEMA-ATEMA-MPC)-SH was tested on MSC culture (Figure 11). Aqueous P(HEMA-ATEMA-MPC) Run 8, P(HEMA-MPC), and MPC solutions were acidic (pH 4.8, 4.2, and 3.8, respectively). It can be assumed, that at high concentrations, these polymers will be toxic due to the low pH. After deprotection of P(HEMAATEMA-MPC) Run 8 with triethylamine and washing with water, a neutral pH was reached (pH 7.6). 2.5 and 5 wt.% of P(HEMA-ATEMA-MPC) Run 8 in culture medium were toxic to the cells, while cell proliferation was unaffected at concentrations < 1 wt.%. In addition, no cytotoxicity was found for P(HEMA-ATEMA-MPC)-SH even at a 5 wt.% concentration, i.e., at the concentration which was toxic for P(HEMA-ATEMA-MPC) Run 8.



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Figure 11. MSCs proliferation in 0, 0.25, 0.5, 1, 2.5, and 5 wt.% of (A) P(HEMA-ATEMAMPC) Run 8 and (B) P(HEMA-ATEMA-MPC)-SH in culture medium measured by the AlamarBlue assay after 1, 3 and 7 days of incubation (mean ± standard deviation, n = 4. ).

CONCLUSION Water-soluble

P(HEMA-ATEMA-MPC)

was

successfully

developed

using

RAFT

polymerization. The indisputable advantage of RAFT polymerization is embodied in the ability to control the molecular weight of the polymer, which can be kept lower than the molecularweight threshold limiting glomerular filtration (~45 kDa). This technique is also suitable for the preparation of block copolymers and various micellar systems, biocompatible antifouling polymer coatings of various surfaces, etc. SEM micrographs confirmed the oriented porosity of 30 ACS Paragon Plus Environment

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the prepared hydrogels, with pore sizes that are suitable for cell accommodation. The solvent regain and degradation of the hydrogels were controlled by changing the concentrations of ATEMA and DTP in the reaction mixture. Both P(HEMA-ATEMA-MPC)-SH and P(HEMAATEMA-MPC) hydrogels proved to be non-toxic. The results suggest that materials can be considered as a promising alternative to poly(ethylene glycol) or polyvinylpyrrolidone for application in tissue engineering and drug delivery systems, e.g., for controlled drug release from subcutaneous implants. ACKNOWLEDGMENT Supported by the Czech Science Foundation (project No. 14-14961S).

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AUTHOR INFORMATION Corresponding Author * Mailing address: Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Heyrovsky Sq. 2, 162 06 Prague 6, Czech Republic. E-mail: [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.


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NOTES We declare no competing financial interests. ¨

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