Electrospun Polythiophene Phenylenes for Tissue Engineering

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Electrospun Polythiophene Phenylenes for Tissue Engineering Eddie Wai Chi Chan, Devasier Bennet, Paul Baek, David Barker, Sanghyo Kim, and Jadranka Travas-Sejdic Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b00341 • Publication Date (Web): 11 Apr 2018 Downloaded from http://pubs.acs.org on April 12, 2018

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Electrospun Polythiophene Phenylenes for Tissue Engineering Eddie Wai Chi Chan†#°, Devasier Bennet,‡,⊥°, Paul Baek†#, David Barker†, Sanghyo Kim‡,§ and Jadranka Travas-Sejdic†#* AUTHOR ADDRESS †

Polymer Electronics Research Centre, School of Chemical Sciences, The University of Auckland, Private Bag 92019, Auckland, New Zealand

#

MacDiarmid Institute for Advanced Materials and Nanotechnology, Victoria University of Wellington, P.O. Box 600, Wellington, New Zealand ‡

Department of Bionanotechnology, Gachon University, Bokjeong-Dong, Sujeong-Gu, Seongnam-Si, Gyeonggi- Do 461-701, Republic of Korea

⊥Noll

Laboratory, Department of Kinesiology, and Huck Institutes of Life Sciences, The Pennsylvania State University, University Park, Pennsylvania, USA

§

Gachon Medical Research Institute, Gil Medical Center, Incheon, 405-760, Republic of Korea

*Corresponding author; Email: [email protected]

° Contributed equally to this work

ACS Paragon Plus Environment

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KEYWORDS: Electrospinning, functionalized conducting polymer, tissue engineering scaffold, cytocompatibility

Abstract: This research focuses on the design of biocompatible materials/scaffold suitable for use for tissue engineering. Porous fiber mats were produced through electrospinning of polythiophene phenylene (PThP) conducting polymers blended with poly(lactide-co-glycolic acid) (PLGA). A peptide containing an arginylglycylaspartic acid (RGD) fragment was synthesized using solid phase peptide synthesis and subsequently grafted onto a PThP polymer using azide-alkyne ‘click’ chemistry. The obtained RGD functionalized PThP was also electrospun into a fiber mat. The electrospun mats’ morphology, roughness and stiffness were studied by means of scanning electron microscopy (SEM) and atomic force microscopy (AFM) and their electroactivity by cyclic voltammetry. The fibers show excellent cytocompatibility in culture assays with human dermal fibroblasts-adult (HDFa) and human epidermal melanocytesadult (HEMa) cells. The electrospun fibers’ roughness and stiffness changed after exposing the fiber mats to the cell culture medium (measured in dry state), but these changes did not affect the cell proliferation. The cytocompatibility of our porous scaffolds was established for their applicability as cell culture scaffolds by means of investigating mitochondrial activity of HDFa and HEMa cells on the scaffolds. The results revealed that the RGD moieties containing PThP scaffolds hold a promise in biomedical applications, including skin tissue engineering.


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Introduction As the field of tissue engineering grows, there is a great need for new material for scaffolds that will provide for more than just structural and mechanical cues.1,2 The application of conducting polymers (CPs) in synthetic tissue engineering scaffolds has been demonstrated to be highly beneficial, as CPs provide means for electrical stimulation of cells enhancing the cells’ growth and proliferation.3–12 For example, Collier et al. have shown that electrically stimulated neuron cells grow almost twice as fast compared to unstimulated cells in in vitro studies.10,13 Application of electrical currents onto CPs may also alter CPs’ physical properties such as charge, stiffness and hydrophobicity, affording “smart”, electrically conducting scaffolds.14–16 The type or function of the cells can be altered via both the structure of the scaffold and the presence of certain biological molecules within the scaffold. Efforts have been also made to add various surface modifications to scaffolds in order to impart physico-chemical and biological cues that promote cell adhesion/proliferation.17,18 The surface topography and mechanical properties are demonstrated to be similarly important and may alter cell behavior.19,20 The interest in electrospun fibers for cell studies and tissue engineering stems from the ability of this fabrication technique to produce highly porous micro- and nano-fiber mats of varying dimensions that mimic the extracellular matrix.21–24 Electrospinning of conducting polymers and their composites has been explored for tissue engineering application,25,26 however with a limited number of CPs utilized27,28 due to their well-known intractability that prevents processing of CPs by common solution-based processing techniques. In general terms, in order to produce CPcontaining fibers, two methodologies are commonly employed: co-spinning of a CP with a carrier polymer, that may also be biocompatible; or with a carrier polymer that is later removed through dissolution.29 In spite of relatively low conductivities of electrospun CP blends with


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biocompatible support polymers, such fibers are still effective in tissue engineering applications, as cells stimulation usually requires relatively low voltages. 30,31 Conducting polymer electrospun tissue engineering scaffolds have been widely employed for electrical cell stimulation.10,13–16 In neural and cardiac tissue engineering applications, electrospun CP fibers mimic the extra cellular matrix and provide electrical stimulus to the cells.32–34 In one of these studies involving cardiac tissue engineering, aligned fibers were used to produce aligned cardiac muscle cells and the conductivity, actuation and anisotropy of the cell scaffold were used to induce synchronized cell beating.27 The use of polythiophene and its derivative, particularly, poly(3,4-ethylenedioxythiophene) (PEDOT) in bioengineering applications has been widely reported in the literature.35–40 Pristine polythiophene was electrospun aided by either crosslinking reactions (magnesium ions as physical cross-linkers), or by utilizing ‘click’ chemistry to add solubilizing side groups onto the CP.41–43 However, a challenge to the prevalent use of PTh and its derivatives is that they are generally difficult to process and of poor mechanical properties.39 We have recently reported synthesis and properties of a range of novel, functionalized thiophene phenylene (ThP) monomers and their polymers (PThP), and discussed advantages of incorporation of functional side groups through the central benzene ring of the monomer on the properties of the obtained polymers, including solubility.44,45 In this study, we utilize these PThP conducting polymers to demonstrate their processability by electrospinning into blend fiber mats with PLGA and PEO. The use of poly(lactic-co-glycolic) acid (PLGA) has attracted considerable interest as a matrix polymer for tissue engineering and in various biomedical applications due to its biocompatibility and the USA Food and Drug Administration (FDA) approval in clinical uses. PThP scaffolds presented here possess multiple functionalities. The PThP backbone has a chemical handle that


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allows the modification with side chains and the insertion of functional groups. As an example we attached the cell adhesive peptide RGD through “click” chemistry. This is one of many potential moieties that can be appended and therefore we demonstrate a general approach to scaffold material with multiple modes of interaction to induce cellular responses. We selected human dermal fibroblast-adult (HDFa) and human epidermal melanocyte-adult (HEMa) cells as these promote human skin tissue formation for regenerative medicine applications. Cytotoxicity of the scaffolds was evaluated and visualized by MTT assay and bio-atomic force microscopy (bio-AFM) imaging. Additional to AFM imaging, biophysical and biomechanical changes of the cells were observed. Overall, this study investigates interactions of the PThP and RGDfunctionalised PThP fiber scaffolds with the cells to provide an insight into the use of this class of conducting polymers electrospun fibers in tissue engineering applications. Experimental Polyethylene oxide (Mn 100,000) (PEO), N,N’-diisopropylcarbodiimide (DIC), 1,1,1,3,3,3hexafluoro-2-propanol (HFIP), trifluoroacetic acid (TFA), triisopropylsilane (TIS), piperidine were obtained from Sigma Aldrich. PLGA (poly(lactic-co-glycolic acid) 75:25) (Mn 110,000 ) was obtained from Purac. (S)-2-Aminopent-4-ynic acid 4 was obtained from Oxchem. 1-Hydroxybenzotriazole hydrate (HOBT), 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexaﬂuorophosphate) (HBTU) were obtained from Sigma Aldrich. PThP polymers of PTMeThP 1 and PTTGThP 2 polymers were prepared using previously reported procedure.44 Synthesis of (S)-2-((((9H-fluoren-9-yl)methoxy)carbonyl)amino) pent-4-ynoic acid 5 To a solution of (S)-2-aminopent-4-ynic acid 4 (1.00 g, 8.85 mmol) in dioxane (24 mL), FmocOSu (3.74 g, 11.1 mmol) and 10% Na2CO3 in H2O (24 mL) was added and the resulting mixture


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was stirred at r.t. for 20 h. Dioxane was removed in vacuo, mixture acidified to pH 3, extracted with ethyl acetate (3 x 10 mL), washed with H2O (20 mL), dried (MgSO4) and solvent removed in vacuo to yield title product 5 (2.76 g, 93%) as a white solid. 1H NMR (400 MHz; DMSO-d6) 2.54-2.69 (2H, m, CβH), 2.90 (1H, t, J = 2.6 Hz, CH), 4.19-4.22 (1H, m, CαH), 4.23-4.25 (1H, CHCH2O), 4.31-4.32 (2H, m, CHCH2O), 7.33 (2H, t, J = 7.4 Hz, Ar-H), 7.42 (2H, t, J = 7.4 Hz, Ar-H), 7.72 (2H, d, J = 7.4 Hz, Ar-H), 7.89 (2H, d, J = 7.4 Hz, Ar-H);

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C NMR (100 MHz;

DMSO-d6) 20.9, 46.6, 52.1, 52.9, 65.8, 73.1, 80.2, 120.1, 125.3, 125.4, 127.1, 127.6, 140.7, 143.7, 155.8, 171.0. The NMR data values are in agreement with literature values.46 Synthesis of RGD containing hexapeptide 6 Wang resin (loading 0.60 mmol/g, 1.97 g, 1.18 mmol) was swelled in 9:1 DMF/CH2Cl2 (30 mL) for 30 min. DMF was drained from the resin and a solution of Fmoc-Gly-OH 7 (1.75 g, 5.90 mmol), HOBT (0.902 mg, 5.90 mmol), DIC (0.743 g, 5.90 mmol), DMAP (0.020 g, 0.118 mmol) and DMF (20 mL) was added and the reaction vessel was shaken for 3 h. The solution was drained from resin. A capping solution of acetic anhydride (0.240 g, 2.36 mmol) and pyridine (0.186 g, 2.36 mmol) was added to the resin and the vessel was agitated for 30 min. The resin was drained and washed with DMF (5 x 10 mL), methanol (10 mL), hexanes (10 mL) and the reaction vessel was dried in desiccator overnight. Fmoc deprotection was achieved by addition of piperidine:DMF (1:4, 15 mL) followed by agitation for 15 min, draining of solution and washed with DMF (5 x 15 mL). The first coupling step was performed by adding HBTU (1.565 g, 4.13 mmol), HOBT (0.631 g, 4.13 mmol), DIPEA (1.23 mL, 7.08 mmol) and FmocAPY 5 (1.185 g, 3.54 mmol) in DMF (10 mL). The solution was agitated for 50 min, drained and washed with DMF (5 x 10 mL). The rest of the peptide chain was built up through consecutive Fmoc deprotection/coupling steps using the following amino acids in the order: Fmoc-Phe-OH 8


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(1.37 g, 0.354 mmol), Fmoc-Asp(OtBu)-OH 9 (1.45 g, 3.54 mmol), Fmoc-Gly-OH 7 (1.04 g, 3.54 mmol) and Fmoc-Arg(PBF)-OH 9 (2.29 g, 3.54 mmol). The resin was then washed with methanol (10 mL), hexanes (10 mL) and CH2Cl2 (3 x 10 mL) and dried in a desiccator overnight. A cleaving solution of 95% TFA, 2.5% H2O, 2.5% TIS (3 mL/100 mg of resin, 20 mL) was then added and the mixture was shaken for 2 h. To the solution of cleaved and deprotected peptide, diethyl ether (10 ml) was added until precipitation occurred. The solid was collected by vacuum filtration, and purified by column chromatography using C18 reverse phase column chromatography to yield title product 8 (513 mg, 66%) as a yellow solid. Mp = 128-132 °C; IR νmax (neat)/cm-1s 3272, 1638, 1534, 1404, 1110; [α]D20 -11 (c 0.5, methanol); 1H NMR (500 MHz, DMSO-d6) 1.44-1.52† (2H, m, Arg(1)-βCH2), 1.55-1.65† (2H, m, Arg(1)-γ), 2.40-2.45† (2H, m, Asp (3)-βCH2), 2.47-2.50† (2H, m, APY(5)-CH2), 2.81-2.85† (2H, m, Phe(4)-βCH2), 3.04-3.09† (2H, m, Arg(1)-δCH2), 3.59-3.67† (2H, m, Gly(6)-αCH2), 3.73-3.79† (2H, m, Gly(2)αCH2), 4.02-4.06† (1H, m Arg(1)-αCH), 4.39-4.44† (1H, m, APY(5)-αCH), 4.44-4.48† (1H, m, APY(5)-δCH), 4.49-4.52† (1H, m Phe(4)-αCH), 4.61-4.65† (1H, m, Asp(3)-αCH), 7.20-7.23 (2H, m, Phe(4)-δCH), 7.41-7.44 (2H, Phe(4)-εCH), 7.58-7.60† (1H, m, Arg(1)-εNH), 7.60-7.65† (2H, m, Arg(1)-NH2), 7.89-7.91 (1H, m, Phe(4)-ζCH), 8.16-8.18† (1H, m, Gly(6)-NH), 8.18-8.22† (1H, m, Phe(4)-NH), 8.23-8.26† (1H, m, Gly(2)-NH), 8.26-8.28† (1H, m APY(5)-NH), 8.588.62† (1H, m, Asp(3)-NH); 13C NMR (125 MHz, DMSO-d6) 25.0 (Arg(1)-βCH2), 28.9 (Arg(1)γCH2), 37.2 (APY(5)-βH2). 37.3 (Arg(1)-δCH2), 37.4 (Phe(4)-βCH2), 37.6 (Asp(3)-βCH2), 40.1 (Gly(6)-αCH2), 40.9 (Gly(2)-αCH2), 49.5 (Asp(3)-αCH), 49.6 (Phe(4)-αCHa), 49.7 (APY(5)δCHa) 51.6 (APY(5)-αCH), 54.3 (Arg(1)-αCH), 80.5 (APY(5)-γC), 125.4 (Phe(4)-ζCH), 127.8 (Phe(4)-εCH), 129.4, (Phe(4)-δCH), 137.6 (Phe(4)-βC), 169.8 (APY(5)-C=O), 170.1 (Phe(4)C=O), 170.4 (Arg(1)-C=Ob), 170.4 (Asp(3)-C=Ob), 170.9 (Gly(6)-C=O), 171.1 (Arg(1)-ζCc),


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171.2 (Gly(2)-C=Oc), 171.5 (Asp(3)-γC=Oc). HRMS (EI) found (MH+) 6462936. C28H40N9O9 requires 646.2944.

†

denotes shifts assigned via 2D NMR correlations,

a-c

denotes assignments

that cannot be definitively discerned from one another. ‘Click’ reaction of hexapeptide 6 with PTMeThP To a solution of azide containing polymers (0.02 mmol) in THF (2 mL), alkyne 6 (0.042 mmol) was added. A solution of ascorbic acid (2.00 µmol) and CuBr(PPh3)3 (2.00 µmol) in water (1 mL) was added dropwise to the polymer solution and the resulting mixture was stirred for 72 h. The mixture was extracted with ethyl acetate (3 x 10 mL) and the combined extracts washed with water (5 mL), brine (5 mL), dried (Na2SO4) and solvent removed in vacuo to give crude triazole, which was washed with water (10 mL) methanol (10 mL) and hexanes (10 mL).

Electrospinning Electrospinning was conducted using Sanitex syringe (1 mL or 5 mL), Thermo Needle (18-gauge x 1 ½ inch). Pump rate was controlled by Adelab Scientific syringe pump, the power was supplied by Bertan High Voltage Power Supply (Series 230) and the collector was a 15 x 15 x 0.1 cm stainless steel collector with or without aluminum foil. SEM and ESEM images were obtained from either FEI (Philips) XL30 S-FEG or FEI Quanta 200 field emission Environmental SEM respectively on platinum coated samples. All PEO matrices nanofiber mats were spun in varying concentrations of chloroform, at 3 mL/h pump rate, 10 cm distance between needle and collector with an applied voltage of 13 kV. All the PLGA matrix nanofiber mats here were spun from the total polymer concentration of 20 w/v% in HFIP pumped at 0.5 mL/h, 10 cm distance between needle and collector and an applied voltage of 13 kV.


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In-vitro cytocompatibility studies The biocompatibility of the nanofibers was investigated by qualitative (optical microscope) and quantitative assays (MTT) using human dermal fibroblast-adult (HDFa) and human epidermal melanocyte-adult (HEMa) cells (purchased from CEFO Ltd, Seoul, South Korea). The HDFa and HEMa cells were cultured using the human fibroblast growth medium (CB-HFGM02), and human melanocyte growth medium (CB-MEL-GM03), respectively. The media was supplemented with respective growth supplements and 1% antibiotics solution and were purchased from CEFO Ltd, Seoul, South Korea. The UV sterilized (irradiation using 8 W Lamp, G30T8, Sankyo Denki, in a biosafety cabinet for 1 hour) electrospun nanofibers (cut to 10 mm in diameter for each sample) were placed in a 6 well plate and the cells (4 × 104 cells/well) were seeded. Culture medium (1 mL) was added 1 h prior to cells seeding to allow for the complete diffusion of culture medium into the fiber structure, and then incubated at 37 °C in 5% CO2 and 95% humidity, followed by the cell culture growth for a week. After the 3rd and 5th days of culture, cell survival was imaged by fluorescence microscopy (Nikon Eclipse Ti, Nikon, Japan) using Nikon NIS-Element software. The mitochondrial activity was quantified using MTT (3[4,5-dimethylthiazol-2-yl]-2,5-diphenyl tetrazolium bromide) proliferation assay kit (Sigma Chemical Co., St. Louis, MO) by following the manufacturer protocol. Cells cultured without nanofibers were considered as the control. The survival of the cells in presence of the fibers was also observed by Calcein-AM (Life Technologies) staining. The actin cytoskeletons were visualized with a fluorescence microscope using rhodamine labeled phalloidin (RLP, for actin cytoskeletons) and 4,6-diamidino-2-phenylindole (DAPI, for nuclei) (Sigma Chemical Co., St. Louis, MO). The morphological characterization of the cells in presence of the nanofibers was observed by field emission scanning electron microscopy (FE-SEM; JEOL-ISM-7500F) and


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high-resolution bio-atomic force microscopy (Bio-AFM; Nanowizard II, JPK instruments, Berlin, Germany) with the inverted optical microscope (Nikon Instruments Eclipse Ti; Amsterdam, Netherlands). Prior to analysis, all cell samples were fixed using 4% paraformaldehyde solution for 30 min. The fixed samples were then dehydrated using ethanol 25%, 50% and 96% for 2 min each, at room temperature and subsequently air-dried.47 Fixation was needed in order to obtain high-resolution images of the cells that were not imaged well in solution; the greater resolution acquired with the fixed cells allowed a further thorough subcellular and substrate examination.48 For FE-SEM examination, the fixed and dried samples were sputter-coated with platinum and visualized using FE-SEM with 15 kV accelerated voltage. Topography, roughness and stiffness of the dried fibers were studied before and after the cell culture with and without cells for all samples using bio-AFM. Conical silicon nitride cantilevers (Si3N4, Au surface; Novascan, Technologies, Inc.) with nominal spring constant of 0.2 N/m were used. Contact mode scans were performed at a rate of 0.2–0.4 Hz, with a high-resolution image quality and a scan size of 100 × 100 µm. For surface roughness analysis, the data obtained from the Bio-AFM height scale images were used to calculate the surface roughness on the dried fibers before and after the cell culture (with no cells region). Using JPK offline data processing software v3.3.25, the root mean square (RMS) roughness (Rq) values were extracted for each sample.49 Force spectroscopy analysis (i.e. nanoindentation), was performed using a soft cantilever (nominal stiffness = 0.01 N/m) with a 5 µm SiO2 particle attached (Novascan, Technologies, Inc.). After selecting the specific fiber regions, the cantilever approached the selected region at a speed of 1 µm/s with a contact force of 1 nN. After the induction of force on the surface, the cantilever was lifted and the deflection curves were recorded to measure the


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relative cell stiffness (Young’s modulus) of the fiber. Young's modulus was calculated using Hertz's contact mechanics model with the JPK data processing software.

Cyclic Voltammetry Cyclic voltammetry experiments were carried out using a CH650 Instruments electrochemical workstation in 0.1 M potassium chloride in water. To perform CVs of the electrospun fibers in solution, the fibers were placed on pre-cleaned indium tin oxide (ITO)-coated glass slides and were held against the substrate by wrapping the fibers on ITO slides with Pt mesh. An approximately 1 cm x 1 cm fiber-coated working electrode window was created by masking the rest of the fibers and ITO with polyimide tape. A multimeter was used to confirm that the Pt mesh was not in contact with the underlying ITO layer, Ag/AgCl wire was used as the reference and Pt wire as the counter electrodes.

Results and Discussion Polythiophene phenylenes (PThPs) possess great versatility in terms of functionalization44,45, as well as good solubility in organic solvents. The PThPs used in this study were PTMeThP 1 and PTTGThP 2; their structures are presented in Figure 1.


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Figure 1: Structure of the PThPs used in electrospinning The PTMeThP 1 and PTTGThP 2 polymers consist of a PThP backbone containing several types of side chains: (i) azide-containing side chains, (ii) a radical polymerization initiating site– containing side chains (for Atom Transfer Radical Polymerization, ATRP) and (iii) either methoxy (PTMeThP 1) or triethylene glycol (PTTGThP 2) side chains. The ratio of the polymers containing these side chains in the polymers was 1:1:3 (as determined by NMR, Figure S1 and S2), respectively and the molecular weights are presented in Table S1. The S1 and S2 differ by the side chain of the ‘diluting’ co-monomer: S1 contains PTMeThP 1 with methoxy side chains, while S2 contains PTTGThP 2 with triethylene glycol side chains. Both S1 and S2 were electrospun to study morphological differences in the fibers formed. Furthermore, ethylene glycol chains are known to prevent non-specific protein adsorption and thus may also have an effect on cell behavior. The ATRP functionality was not utilized in this work to reduce


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the complexity of the cell biocompatibility studies; however, we incorporated these as we have previously shown these can be readily utilized in grafting of various types of polymeric side chains44 that may have an important use in tissue engineering applications, such as anti-fouling.9

Initial attempts on electrospinning of PTMeThP 1 and PTTGThP 2, were undertaken with polyethylene oxide (PEO) as a carrier polymer. PEO is commonly electrospun polymer that gives fibers of a consistent size50 and is a well-studied in electrospinning.51–54 It was employed in the preliminary experiments to assess how the carrier polymer concentration and overall polymer concentration effects the morphology of the fibers. To alter the composition of the nanofiber mats the concentrations of the polymers in the solution were varied. The SEM/ESEM images of the prepared PEO-PThP fibers are presented in Figure S3-12. Table S2 provides different weight ratios of PThP to PEO utilized, total polymer concentrations in solution and the average diameter of the obtained fibers. These results showed that the PThP polymers can be coelectrospun to form well-blended nanofiber composites with the fiber diameter modulated by the electrospinning conditions. However, the cell medium dissolved PEO in PThP/PEO fiber mats and the PThP component was unable to retain the fiber structure. Therefore, the following electrospinning experiments were performed using biocompatible, but water insoluble, PLGA as a carrier polymer matrix.

The use of PLGA in tissue engineering is well documented with applications in clinical settings.55,56 Here, hexafluoroisopropanol (HFIP) was used as solvent for the electrospinning,57 rather than chloroform, as it produced well-defined and uniform fibers, likely due to the increased solubility of PThP in the higher polarity solvent. Conditions of 13 kV electric field, a


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10 cm needle-collector distance and a pump rate of 3 mL/h produced excellent fiber mats. PTMeThP 1 and PTTGThP 2 were electrospun with PLGA, at the ratio of 1:4 (w/w), from a 20 w/v% total polymer concentration solution in HFIP, to obtain the fiber mats of PMeThP-PLGA (P1) and PTTGThP-PLGA (P2). Figure 2 and Figure 3 present the SEM and AFM images of the PTMeThP-PLGA P1 and PTTGThP-PLGA P2 mats, respectively. These mats were easy to peel off from the metal collector and had fabric-like properties, making them easy to handle. The fiber mats appear porous throughout the whole mat thickness, but interestingly the fibers’ diameters were very different, in the range of 1.27 ± 0.17 µm and 0.30 ± 0.05 µm for P1 and P2, respectively, likely due to the higher viscosity of the polymer solution of P1 relative to P2.


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Figure 2: Electrospun fibers of PTMeThP-PLGA P1, consisting of PTMeThP 1 (20 w/w%) and PLGA (80 w/w%). Top row: SEM images of: (A) Low magnification (bar 20 µm), (B) Front side and (C) Back side of the mat (bar 5 µm). Bottom row: (D) AFM 3D height image and (E) a height profile of fibers along the 50 µm line horizontally.


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Figure 3: Electrospun fibers of PTTGThP-PLGA P2, consisting of PTTGThP 2 (20 w/w%) and PLGA (80 w/w%). Top row: SEM images: (A) Low magnification (bar 20 µm), (B) Front side and (C) Back side (bar 5 µm). Bottom row: (D) AFM 3D height image and (E) a height profile of fibers along the 50 µm line horizontally.

To demonstrate the use of the functional moieties on PTMeThP 1 for further functionalization with biomolecules, that may direct or stir cell behavior on these scaffolds, a hexapeptide 5 containing an RGD motif was ‘clicked’ on PTMeThP 1. RGD peptides are cell adhesive which enhances cell attachment through interaction with integrin on cells.58–60 The RGD peptide 5 was synthesized using an unnatural amino acid 3 containing an alkyne side chain, which was then Fmoc protected, giving 4, and the hexapeptide 5 was synthesized using solid phase peptide synthesis. Peptide 5 was then ‘clicked’ on to PTMeThP 1 to obtain PMeThP-RGD 6 (Scheme 1).


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Scheme 1: Synthesis of PTMeThP-RGD 6 i) Fmoc-OSu, Na2CO3, dioxane, r.t., 20 h, 93%; ii) Resin Loading: Wang Resin, Fmoc-Gly-OH, HOBT, DIC, HOBT, DMAP, DMF, rt, 3 h; FmocDeprotection: piperidine, DMF, rt, 30 min; Peptide Coupling: HBTU, DIPEA, DMF, rt, 1 h; Peptide cleavage: TFA, H2O, TIS; 66% (6 steps), iii) PTMeThP 1, CuBr(PPh3)3, THF, rt, 72 h, 40%. The resultant polymer 6 was co-electrospun with PLGA under the same conditions as the above-discussed PTMeThP-PLGA P1 and PTTGThP-PLGA P2. Figure 4 A-C present the SEM images of the PMeThP-RGD-PLGA fibers (P3) that alongside the AFM topography image and a height profile (Figure 4 D, E), show the uniformity in the size of the fibers that spun without


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beading, as well as the good porosity of the mat. The PTMeThP-RGD-PLGA (P3) electrospun fibers had a consistent diameter of 0.37 ± 0.10 µm, with straight, tubular morphology. The fibers are considerably thinner than P1 despite the similar backbone used and of similar diameter to P2, likely due to the increased polarity of the P2 and P3 polymers due to ethylene glycol and RGD moieties.

Figure 4: Electrospun fibers PTMeThP-RGD-PLGA P3, consisting of PTMeThP-RGD (20 w/w%) and PLGA (80 w/w%). Top row: SEM images of: (A) Low magnification (bar 100 µm) (B) Front side and (C) Back side (bar 5 µm) of the mat. Bottom row: (D) AFM 3D height image and (E) a height profile of fibers along the 50 µm line horizontally.


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A pure PLGA fiber mat was also prepared using the same conditions as above, for comparison purposes, especially in cytocompatibility studies (where it is referred to as Control, see below). The SEM images and Bio-AFM studies are presented in the supporting information (Figure S13). The fiber diameters are 1.25 ± 0.17 µm, which is similar to P1 but thicker than P2 and P3.

In-vitro cytocompatibility studies were performed on all fibers - P1, P2, P3 and Control to test their potential as scaffold materials. The HDFa and HEMa cells on the nanofiber mats were characterized by FE-SEM, Bio-AFM and by fluorescence microscopy to study the effect of morphology, topography, surface roughness and stiffness of the fibers, which are known to affect the cell attachment and proliferation.61 SEM images of HDFa cells (Figure 5) and HEMa cells (Figure 6), grown for 3 and 7 days of cell culture, on PLGA Control fiber mat, is shown in Figures 5A and 6A, on PTMeThP-PLGA P1 fiber mat in Figures 5B and 6B, on PTTGThPPLGA P2 fiber mat in Figures 5C and 6C, and on PTMeThP-RGD-PLGA P3 fiber mat in Figures 5D and 6D. SEM images obtained on different locations on these substrates are presented in the Supporting Information (Figure S14 and S15).


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Figure 5: SEM images of HDFa growth on electrospun fiber mat after 3 and 7 days of cell culture. A: PLGA (Control); B: PTMeThP-PLGA P1; C: PTTGThP-PLGA P2; D: PTMeThPRGD-PLGA fiber mat P3. Scale bars are given in image.


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Figure 6: SEM images of HEMa growth on electrospun fiber mat after 3 and 7 days of cell culture. A: PLGA (Control); B: PTMeThP-PLGA P1; C: PTTGThP-PLGA P2; D: PTMeThPRGD-PLGA P3 fiber mats. Scale bar in all SEM images is 10 µm.


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Both types of the cells, on all of the fiber mats, showed attachment and proliferation, as seen in Figures 5 and 6. The cell proliferation on P1, P2 and P3 were comparable to the PLGA Control fiber mat. Images of HDFa and HEMa cell cultures on all investigated fibers clearly show cytocompatibility of the polymer fibers with the cell growth. There is increased cell attachment with the increase in cell culture cultivation time from the 3rd to the 7th day, without any noticeable differences of the cells’ growth on the fibers containing PTMeThP P1 to that on the PLGA fiber mat. The cells appear to mainly adhere onto the surface of the fibers, with few a possibly protruding into the fiber mats. The attached cells’ actin filaments penetrate and fibrillary extend along the fibers. The PTMeThP-RGD-PLGA P3 fiber mats showed similar results with no obvious differences in the HDFa and HEMa cells’ growth compared to the fibers without the RGD moiety attached. As the cells adhere well to the fibers anyway, the beneficial effect of enhanced cell attachment through the RGD moiety is not obvious in this instance. However, more extensive filopodia of HEMa cells (Figure 6D) - which are responsible for cell attachment, organization of the cytoskeleton and proliferation on the cells62 - were observed on PTMeThPRGD-PLGA P3 fiber mats, indicating an improved cell migration and HEMa cell–cell interactions. This observation was reflected in the cell-based assays for cell proliferation, Figure 7. The proliferation efficiency of the cultured cells on the various fibers, at the two incubation periods, were calculated by evaluating mitochondrial activity through the use of a cell proliferation reagent MTT kit, as shown in Figure 7A. When comparing the three substrates, HEMa cells on P3 fiber showed significant enhancement in proliferation behavior during the 3 and 7 days period, which may be due to the present RGD binding moieties. The fluorescence of the metabolically active cells on fibers was visualized using Calcein-AM staining (Figure 7B). In particular, we noticed that the greatest numbers of both cell types, HDFa and HEMa, were


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attached to the P3 fibers containing the RGD binding moieties. Particularly, the HDFa cells are growing into P3 fiber scaffolds, but not into the P2, which may be due to the RGD functionality being dispersed within the fibers. Moreover, the MTT and Calcein AM cell viability assays proved that the RGD peptides enhance the cell attachment and proliferation. Having RGD peptides covalently attached to the conducting polymer is considered advantageous in contrast to simply incorporating the RGD peptides into the electrospinning solution, as the later may lead to leaching out of the peptides and therefore decreased effect on the cell proliferation.

Figure 7: Cell viability assay: (A) MTT cell proliferation assay demonstrating mitochondrial activity of HDFa and HEMa cells after 3rd and 7th day of culture on fiber mats. Results are given as (mean ± SD) of 3 independent experiments. Bars show statistically significant differences (P>0.05). (B), Calcein AM cell viability assay. Calcin AM is used for labeling live cells of HDFa and HEMa cells after 7th day of culture on fiber mats.

Additionally, the cellular behavior, and therefore biocompatibility, was assessed through a cell viability assay, where phalloidin and DAPI were used to stain the actin filament and cell nuclei, respectively. The results are shown in Figure 8. After 7 days of cell culture, the cells exhibited


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higher cell density on the P3 fiber mat than on the P1, P2 and Control, indicating the increased cells population. The phalloidin staining showed that HEMa cells grown on P3 fiber retain the extended and stronger actin filaments than on other fiber mats. Moreover, HEMa cells demonstrated relatively higher cellular survival and proliferation compared to the HDFa cells, possibly due to the difference in the surface topography and roughness of the P3 fiber matrix on these cells.63,64

Figure 8: Fluorescence micrographs of DAPI and rhodamine-phalloidin staining of (A) HDFa and (B) HEMa cells grown for 7 days on fiber mats (Control, P1-P3). Figure 9 shows the height images of the fibers-coated cells (HDFa (A) and HEMa (B) cells), captured by bioAFM imaging after 7 days culturing on the fiber mats (Control, P1, P2 and P3,


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and on bare substrate). The cells appear to be distributed over the substrates’ surface. Both cells exhibited their characteristic shape, and, importantly, the appearance of the actin-rich membrane protein protrusions along the fibers, which contribute to invasion through the pores of scaffolds (indicated by the arrows). Figure 9 clearly shows the cell attachment, actin filament penetration and predominant fibrillar extensions onto P3 fibers, which we suggest to be likely related to RGD moieties presented on the fibers. The topographies observed in Figure 9 correlated well with the cytotoxicity data. In addition to topography, surface roughness and stiffness are equally important for various cellular processes, including cell attachment, proliferation and migration.65 Figure 10A and B (top) show the changes in the surface roughness and stiffness of the fixed cells, respectively, on the fiber mats, determined by examining a 10 × 10 µm area (Figure 9, marked with a gray square bracket in the vertical deflection image) using AFM and JPK data processing software applied to the images. The mean surface roughness values of the cells on the Bare substrate indicated that these cells were relatively smooth. The roughness of the cells on the fiber mats resulted in minor increment in roughness. Similarly, the stiffness for the fixed cells on the Bare substrate was found to be 7 ±1.4 kPa, and for P1-P3 and Control values were between 14 ±4.5 kPa to 20 ±4.9 kPa. The stiffness values of the cells on all of the samples were comparable to the stiffness of human skin cells and fibrotic tissues.66,67 Overall, the cells grown on the fiber mats show higher stiffness and roughness compared to the bare substrate (Figure 10 A and B); still the cell’s spreading area and proliferation was higher on the fiber mats (Figure 7).

Additionally, the surface roughness of the fibers measured in dry state before and after immersing in the cell culture for 7 days (without the cell areas), was estimated. The root mean


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square roughness (Rq) of individual fibers is shown in Figure 10C. The wetting of the fibers showed an effect on the surface roughness of all fibers. The surface roughness and fiber diameter are important factors affecting the surface properties. Modulating surface roughness of 3D scaffolds was shown to be a significant factor in scaffold design.68 During the 7 days of the cells culture, the roughness of the fiber decreased for all fibers; however, in this study, those changes in roughness did not affect the cell proliferation, nor there was a noticeable change observed in cell attachment and viability (Figure 7). A somewhat higher roughness was observed for the P2 and P3 fibers, which also showed higher cell proliferation (Figure 7A). The higher cell proliferations in P3 fiber is also likely related to RGD moieties.69


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Figure 9. Bio-AFM images showing topographical changes (vertical deflection) in cells grown on different fiber mats. Bio-AFM images of HDFa cells (left panel, A) and HEMa cells (right panel, B) on (from top to bottom) PLGA (Control), PTMeThP-PLGA P1, PTTGThP-PLGA P2, PTMeThP-RGD-PLGA P3 fiber mats and Bare substrate. Images of the cells on the Control show relatively smooth cells’ surfaces. The cells attached on the fiber mats show outgrowth, suggesting increased cell–substrate interactions and migration on the fibers. Scan size = 100 × 100 µm for all images.

The stiffness of the fibers, before and after cell culture (carried out on the dried mats), was measured using AFM nanoindentation technique (Figure 10D). The average stiffness for Control PLGA fibers before the cell culture was 746 ±16.4 kPa, similar to the 752 ±16.4 kPa for P3 fibers and 806 ±20.5 kPa for P1 the fibers, whereas P2 fibers had slightly lower stiffness of 551 ±30.4 kPa. The measured values are comparable to the stiffness of other tissue engineering materials.70 After 7 days of cell culture the stiffness of the fibers in dry state significantly decreased for all fiber types to the range 97 – 110 kPa, comparable to the stiffness of in-vivo collagen fibers (109.8 ±13.2 kPa).71 Such decreases in the fibers’ stiffness are suggested to originate from the protein adsorption from the culture media as well as the extracellular proteins from the cells into the fibers. However, these changes did not affect the cell growth and proliferation. Overall, the performed cell assays studies suggest that all of the investigated scaffolds are biocompatible; however, detailed mechanisms of interactions between cells and scaffolds may need further studies. The non-toxicity of the PThP-PLGA fibers is similar to other electrospun polythiophene fibers reported in the literature.72–76 This indicates that the inclusions of sidechains


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and the benzene moiety in the conjugated polymer backbone have no effect on the toxicity of PThP.

Figure 10: Surface roughness and Young’s modulus of cells (A and B), and individual fibers before and after cell culture (C and D). (A), The root mean square roughness (Rq); (B) Young’s modulus measured in the cytoplasmic regions for the corresponding samples of PLGA (Control), PTMeThP-PLGA P1, PTTGThP-PLGA P2, PTMeThP-RGD-PLGA P3 fiber mats and HEMa and HDFa cell (Bare) substrate. Values are expressed from the 10 × 10 µm scale area (gray square bracket) of cells from the deflection images (Figure 9). C), Changes in the roughness of the individual fibers before and after 7 days cell culture. (D) Changes in the


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stiffness of the investigated electrospun fibers before and after 7 days cell culture. The scanning area for roughness and stiffness analysis was on the fibers regions. Data are represented as means ± SD. The electrochemical characterization of the PTMeThP-PLGA fiber mats showed mainly a capacitive behavior of the mats and not well-pronounced oxidation/reduction wave of the PThP (Figure S16-S18). This is not unexpected, as the proportion of PTMeThP (20 wt%) in the total fiber was low. Results of the preliminary study on incorporating larger proportion of conjugated polymer in the fiber blends are given in the Supporting Information (Figure S19). The voltammograms obtained at different scan rates still demonstrate capacitive behavior, but with clearly seen electrochemical redox wave of the PThP backbone, with oxidation and reduction peaks found at 0.29 V and 0.25 V (vs. Ag/AgCl wire), respectively. This result signifies that with further optimization of the fiber’s composition the electrochemical properties of the fiber mats could be enhanced, which would be desirable for efficient stimulation of cells.

Conclusion We report, for the first time, electrospinning of the PThP class of conducting polymers, using PLGA and PEO as carrier polymers. The azide moiety on the polymer provides a convenient and easily accessible site for further functionalization of the polymer with biomolecules that can provide biological cues to the cells to direct and stir their behavior. We have demonstrated the principle by incorporating an RGD peptide using ‘click’ chemistry. An ATRP functional site on these polymers provides another convenient route to adding desired properties to the scaffolds. All of the investigated variants of the PThP polymer electrospun well into the fibers without


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beads. The growth of the cells studied, HDFa and HEMa, was not hindered by the presence of the PThP. Various cell assays collectively confirmed that all of the investigated scaffolds are cytocompatible, demonstrating the polymers’ biocompatibility. The fibers with RGD moieties showed a significant enhancement in the cells’ proliferation, especially at longer cell culture times. Electrochemical activity of the PThP in the PThP-PLGA composite fiber is maintained, albeit the redox processes of the PThP in the fibers are not well pronounced. Further studies will seek to optimize the fibers’ composition regarding their electrochemical properties, as well as to investigate the stimulation of electrically responding cells, such as neurons, and functionalization with other biological cues, such as growth factors. The novel electrospun scaffolds based on PThP described in this study showed a potential for use in biomedical applications, including tissue engineering applications.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge online. 1

H NMR and GPC traces of PTMeThP 1 and PTTGThP 2 (Figure S1 and S2). Table S1

presents the molecular weight of PTMeThP 1 and PTTGThP 2. SEM/ESEM images of PEOPThP fibers (Figure S3-12), electrospinning and diameters of PEO-PThP fibers (Table S2). SEM and AFM studies on PLGA nanofibers (control) (Figure S13). Low magnification SEM images of HDFa and HEMa cell growth (Figure S14 and S15). Cyclic voltammetry study of pure ITO coated glass slide, P1 and P3 (Figure S16-18). A preliminary study on incorporating larger proportion of conjugated polymer in the fiber blends (Figure S19).

AUTHOR INFORMATION


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Corresponding Author E-mail: [email protected] Author Contributions The manuscript was written through contributions from all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest. ACKNOWLEDGMENT This work was supported by the Royal Society of New Zealand, under a New Zealand - Korea Joint Research Projects grant. REFERENCES (1)

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For Table of Contents use only Graphical Abstract

Electrospun Polythiophene Phenylenes for Tissue Engineering Eddie Wai Chi Chan, Devasier Bennet, Paul Baek, David Barker, Sanghyo Kim and Jadranka Travas-Sejdic*


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Electrospun Polythiophene Phenylenes for Tissue Engineering

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