Article pubs.acs.org/Biomac
Lysine-Appended Polydiacetylene Scaffolds for Human Mesenchymal Stem Cells V. Haridas,*,† Sandhya Sadanandan,† Pierre-Yves Collart-Dutilleul,‡ Stan Gronthos,§ and Nicolas H. Voelcker*,∥ †
Department of Chemistry, Indian Institute of Technology Delhi, New Delhi-110016, India Bio-Nano Laboratory, Faculty of Dentistry, Montpellier University, Montpellier, France § Centre for Stem Cell Research, Robinson Institute, North Adelaide, SA-5006, Australia ∥ Mawson Institute, University of South Australia, Adelaide, SA-5001, Australia ‡
S Supporting Information *
ABSTRACT: We report on the self-assembly based fabrication of fibrous polymers for tissue engineering applications. Directed self-assembly followed by polymerization of lysineappended diacetylenes generated a variety of polymers (P1− P5) with distinct chemical properties. The self-assembly along with the conjugated double and triple bonds and rigid geometry of diacetylene backbone imposed a nanofibrous morphology on the resulting polymers. Chemical properties including wettability of the polymers were tuned by using lysine (Lys) with orthogonal protecting groups (Boc and Fmoc). These Lys-appended polydiacetylene scaffolds were compared in terms of their efficiency toward human mesenchymal stem cells adhesion and spreading. Interestingly, polymer P4 containing Lys Nα-NH2 and Lys Nε-Boc with balanced wettability supported cell adhesion better than the more hydrophobic polymer P2 with Nε-Boc and Nα-Fmoc or more hydrophilic polymer P5 containing free Nε and Nα amino groups. The molecular level control in the fabrication of nanofibrous polymers compared with other existing methods for the generation of fibrous polymers is the hallmark of this work.
1. INTRODUCTION Tissue engineering (TE) and regenerative medicine are promising avenues for the treatment of various human ailments. TE aims at recreating tissues that are defective or lost. An important aspect of TE is the seeding of mammalian cells on natural or synthetic extracellular substrates to create a 3D tissue.1 Scaffold plays a critical role in tissue regeneration and repair;2 therefore, various natural macromolecules including silk,3 chitosan,4 and collagen5 are explored for TE applications. Synthetic polymers such as poly(ethylene glycol) (PEG), poly(vinylalcohol) (PVA), poly(acrylic acid) (PAA), polyglycolic acid (PGA), polylactic acid (PLA), polycaprolactones (PCL), poly(propylenefumarate) (PPF) have been investigated as scaffold materials.6 These homopolymers and their copolymers are examples of degradable polymers, but their hydrophobic nature leads to poor wetting, reducing the cell attachment without prior surface modification.7 In order to overcome these disadvantages, hydrogels are extensively used in TE applications.8 Hydrogels are three-dimensional networks of hydrophilic polymers that can absorb water without undergoing dissolution of the polymer and allow the transport of low molecular weight solutes and nutrients into the hydrogel as well as the cellular waste out of the hydrogel, which are critical to cellular growth.9 © 2013 American Chemical Society
Synthetic hydrogels are typically not degradable without incorporation of degradable peptide cross-links and neither do hydrogels provide cues for cell adhesion without incorporation of cell adhesion mediating peptides. Therefore, many synthetic polymers incorporating Arg-Gly-Asp (RGD) peptides, which are found in proteins of extra cellular matrix (ECM), were synthesized and have been investigated for cell adhesion and growth. RGD peptides are known to mimic cell adhesion proteins and bind to integrins, thus scaffolds containing these peptides act as promising TE scaffolds.10 Natural and synthetic polymeric scaffolds face serious limitations such as difficulty in controlling their pore size, shape, and fiber diameter favorable for tissue growth.11 Approaches like electrospinning and phase separation are used for synthesizing fibrous and porous materials for TE, but these methods have no molecular level control on the morphology of the polymers. The ability to control the nanoand microstructures of the scaffold material requires a molecular level approach that relies on intermolecular interactions. Stupp and co-workers designed and synthesized Received: November 4, 2013 Revised: December 17, 2013 Published: December 23, 2013 582
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Figure 1. General representation of self-assembly and topochemical polymerization of diacetylenes.
peptide amphiphiles (PAs) for mimicking collagen-like fibers. PAs were also investigated for the TE applications.12 To fabricate polymeric organic materials with tailored functions and desirable surface morphology requires an approach that relies on precise arrangement of the carefully selected component molecules. The precise arrangement of molecules is achieved only by noncovalent interactions. One of the approaches is the organization of the molecules in a regular array using careful design driven by hydrogen bonding interactions. In order to fix the resulting self-assembled structures and avoid premature disassembly, topochemical diacetylene polymerization can be contemplated (Figure 1). Diacetylenes are a class of compounds which contain conjugated triple bonded systems which can undergo UV polymerization (λ = 254 nm) to yield very stable polydiacetylenes (PDAs). The diacetylene linkage confers rigidity, and the right choice of appended units add functional properties to the materials.13 Frauenrath and colleagues demonstrated the utility of this supramolecular approach to fabricate nanofibrils using conjugated systems.14 Supramolecular self-assembly has also proven to be a versatile tool for formation of organo gels.15
hexane/ethyl acetate. Melting points were recorded in a Fisher-Johns melting point apparatus and were uncorrected. Optical rotations were measured using a Rudolph Research Analytical Autopol V Polarimeter; concentrations are given in grams/100 mL. IR spectra were recorded on a Nicolet, Protégé 460 spectrometer as KBr pellets. 1H and 13C NMR spectra were recorded on a Bruker-DPX-300 (1H, 300 MHz; 13 C, 75 MHz) spectrometer using tetramethylsilane (1H) as an internal standard. Coupling constants are in Hz and the 1H NMR data are reported as s (singlet), d (doublet), br (broad), br d (broad doublet), t (triplet), and m (multiplet). HRMS were recorded with micrOTOF-Q II using ESI technique. Polymerization reactions were performed by irradiating under a high intensity mercury lamp (OmniCure Series 1000). Atomic Force Microscopy (AFM). AFM imaging and measurements were performed using a Bioscope Catalyst AFM (Bruker Corporation, Billerica, MA) with silicon probes. The sample (10 μL) was dropcasted on a freshly peeled mica surface followed by drying under nitrogen flow. The standard tapping mode was used to image the morphology at room temperature in air. A spring constant of 20−80 N/m was used. A single third order flattening of height images with a low pass filter was done followed by section analysis to determine the dimensions of the images. Scanning Electron Microscopy (SEM). SEM images were recorded using ZEISS EVO Series Scanning Electron Microscope EVO 50 operating at an accelerating voltage of 0.2−30 kV. For SEM, a 10 μL aliquot of the sample solution was drop-casted on a glass coverslip, dried, and coated with ∼10 nm of gold. High-Resolution Transmission Electron Microscopy (HRTEM). Images were recorded on a Tecnai G2 20 electron microscope operated at an accelerating voltage of 200 kV. Samples were prepared by drop-casting the sample solutions (P1 in chloroform and P3 in acetonitrile) on 200 square mesh carbon-coated copper grids. Contact Angle Measurements. The water contact angles were measured using Kruss goniometer drop shape analysis system (DSA
2. MATERIALS AND METHODS 2.1. Materials. Reagents were purchased from Chem Impex International, Sigma-Aldrich, and Alfa Aesar. Reagents were used without further purification. All solvents employed in the reactions were distilled or dried using appropriate drying agents prior to use. Reactions were monitored by thin layer chromatography (TLC). Column chromatography was performed on silica gel (100−200 mesh) columns, which were generally made from slurry in hexane, 583
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Preparation of P3. To a solution of P2 (50 mg) in dry dichloromethane (1 mL) was added 0.5 mL of TFA and stirred for 3 h. Reaction mixture was evaporated under high vacuum to yield 23 mg of P3. Preparation of P4. To a solution of P2 (100 mg) in DMF (1.4 mL) was added piperidine (0.6 mL) and stirred at RT for 2 h. The solution was washed with hexane and evaporation of DMF afforded 42 mg of P4. Preparation of P5. To a solution of polymeric gel P1 (50 mg) in 2 mL dichloromethane was added 1 mL of trifluoroacetic acid and stirred for 4 h. Reaction mixture was evaporated under high vacuum to yield P5. Cell Attachment, Viability, Morphology, and Spreading. DPSC were seeded onto the polymers P1, P2, P3, P4, and P5 at a cell density of 5 × 104 cell/mL. Glass coverslips were used as controls. Cultures were incubated for 4, 24, 48, and 72 h at 37 °C with 5% CO2, in a humidified incubator. To determine cellular viability and morphology on the surfaces, cells were stained with 50 μg/mL of fluorescein diacetate (FDA) and 2 μmol/mL of propidium iodide (PI) and incubated for 3 min at 37 °C. Samples were subsequently washed with phosphate buffered saline (PBS) to remove unbound dye. It was then analyzed under the fluorescence microscope at excitation wavelengths of 480 nm for FDA and 630 nm for PI. Images were captured on a Nikon Eclipse 50i epifluorescence microscope. For cell counts, samples were fixed with 2.5% glutaraldehyde for 1 h and stained with 50 μg/mL of Hoechst 33342 for 30 min to stain the nuclei. Samples were then washed with PBS buffer to remove any nonadherent cells and unbound dye and observed under the fluorescence microscope at an excitation wavelength of 360 nm. Cell counts were conducted at five different locations on the surface of each sample (four peripheral and one central) in areas measuring 1400 × 1050 μm. Statistical differences between polymers were determined using the Student’s t test with p < 0.05 deemed as statistically significant. To evaluate cell morphology and spreading, DPSC were stained for actin and nuclei with phalloidin and Hoechst 33342, respectively. Cells were seeded and incubated on different polymers as previously described. After removing the culture medium, cells were rinsed thrice with PBS, fixed with 4% paraformaldehyde for 20 min, and then washed thrice to remove excess paraformaldehyde. DPSC were permeabilized with 0.5% Triton X-100 in PBS at 4 °C for 15 min. Samples were incubated for 1h with TRITC-labeled phalloidin (1:200) at 37 °C in the dark, then incubated with Hoechst 33342 for 10 min at room temperature, and washed twice with deionized water. Samples were observed under fluorescence microscope at an excitation wavelength of 360 nm for Hoechst staining and 630 nm for phalloidin, respectively. Osteogenic Differentiation Assay. DPSC were incubated on polymers P1, P2, P3, P4 and glass coverslip (control) for 3 and 7 days in osteogenic medium (αMEM supplemented with 15% fetal calf serum, 100 μM L-ascorbic acid, 2 mM L-glutamine, dexamethasone 0.1 μM, KH2PO4 2.6 mM, 100 IU/mL penicillin, and 100 μg/mL streptomycin). Cell differentiation was assessed by fluorescence staining of alkaline phosphatase (ALP) and nuclei. In order to stain for ALP, samples were washed with PBS and fixed with 4% paraformaldehyde for 20 min at room temperature. Cells were permealized with 0.5% Triton X-100 in PBS for 15 min at 4 °C, then blocked with 1% BSA in PBS for 1h at room temperature. Samples were incubated with rabbit ALP antibodies (1:100 in 1% BSA) at 4 °C overnight. After three times of rinsing with PBS, samples were incubated with rat antirabbit secondary antibodies (1:200 in 1% BSA, PE-conjugated rat antirabbit IgG) for 1 h at room temperature. Finally, cell nuclei were stained with 50 μg/mL Hoechst 33342 for 30 min at room temperature. Samples were observed under the fluorescence microscope at an excitation wavelength of 360 nm for Hoechst staining and 488 nm for ALP staining.
100) at ambient temperature. Average water contact angles were obtained by measuring the same sample at five different positions. Gel Permeation Chromatography (GPC). Molecular weight distribution was analyzed by Waters HPLC 1525 (based on polystyrene calibration) equipped with a Styragel HR4 THF 7.8 × 300 mm column using refractive index detector. Human Dental Pulp Stem Cells (DPSC). Mesenchymal stem cells derived from human teeth (dental pulp stem cells, DPSC) were isolated as previously described.16 Briefly, discarded normal human impacted third molars were collected with informed consent of patients undergoing routine extractions at the Dental Clinic of the University of Adelaide, under approved guidelines set by the University of Adelaide and Institute of Medical and Veterinary Science Human Subjects Research Committees. Tooth surfaces were cleaned and cracked open to reveal the pulp chamber. The pulp tissue was gently separated from the crown and root and then digested in a solution of 3 mg/mL collagenase type I and 4 mg/mL Dispase for 1 h at 37 °C. Single-cell suspensions were obtained by passing the cells through a 70 μm strainer. Cultures were established by seeding singlecell suspensions (1−2 × 105) of dental pulp into T-25 flasks in growth media, (αMEM supplemented with 10% fetal calf serum, 100 μM Lascorbic acid, 2 mM L-glutamine, 100 IU/mL penicillin, and 100 μg/ mL streptomycin), then incubated at 37 °C in 5% CO2. 2.2. Methods. Preparation of Alkyne Derivative (1). To an icecooled solution of Bis(Boc)Lysine (1.0 g, 2.89 mmol) in 100 mL of dry dichloromethane was added N-hydroxysuccinimide (0.330 g, 2.86 mmol) and dicyclohexylcarbidimide (DCC; 0.590 g, 2.86 mmol) and stirred for 5 min. To this stirring solution was added propargylamine (0.18 mL, 2.869 mmol) and stirred for 24 h. The reaction mixture was filtered and washed with 0.2 N H2SO4, saturated NaHCO3, and water. The organic part was dried over anhydrous Na2SO4 and concentrated in vacuum to yield 0.942 g of the product.15c Preparation of D2. To a solution of 1 (0.100 g, 0.26 mmol) in 30 mL of acetonitrile was added Hay catalyst (a well-stirred solution of CuCl (10 mg, 0.10 mmol) and TMEDA (0.033 mL, 0.22 mmol) in 5 mL of acetonitrile for 5 min) and stirred under air. After 3 h, more Hay catalyst (5 mL) was added and left to stir for 12 h. The reaction mixture was evaporated, redissolved in dichloromethane, and washed sequentially with 0.5 N aqueous H2SO4, water, saturated aqueous NaHCO3 solution, aqueous NH4Cl + NH4OH (9:1) solution, and water. The organic part was separated, dried over anhydrous Na2SO4, filtered, and evaporated to yield 0.072 g of D1.15c Preparation of Alkyne Derivative (2). To an ice-cooled solution of Fmoc-(Nε-Boc)-lysine (0.500g, 1.067 mmol) in 100 mL of dry dichloromethane was added N-hydroxysuccinimide (0.135 g, 1.173 mmol) and dicyclohexylcarbodimide (0.242 g, 1.173 mmol) and stirred for 5 min. To this solution was added propargylamine (0.075 mL, 1.173 mmol) and this was stirred for 24 h. The reaction mixture was filtered, washed the filtrate with 0.2 N H2SO4, saturated NaHCO3, and water. The organic part was dried over anhydrous Na2SO4 and concentrated in vacuum to obtain 0.448 g of the product. Preparation of D2. To a solution of 2 (0.300 g, 0.593 mmol) in 100 mL of acetonitrile was added Hay catalyst (a well-stirred solution of CuCl (25 mg, 0.257 mmol) and TMEDA). Reaction was monitored using TLC. Hay catalyst was added again and again until the starting material was completely gone. The reaction mixture was filtered, the residue obtained was washed with NH4Cl/NH4OH (9:1), 0.2 N H2SO4, saturated NaHCO3, and finally with water. It was further dried to obtain 0.232 g of D2. Preparation of P1. Compound D1 (20 mg) was dissolved in 0.4 mL of ethyl acetate and to this was added 0.6 mL of hexane to obtain the D1 gel. This gel was irradiated under a high intensity mercury lamp (OmniCure Series 1000) for 3 min to obtain the polymeric gel. It was carefully transferred to the glass coverslip for the cell adhesion analysis. Preparation of P2. Polymer P2 was prepared by irradiation of D2 under a Mercury lamp for 3 min. A total of 20 mg of P2 was dissolved in 1 mL of tetrahydrofuran and sonicated for 2 min. The polymeric solution of P2 was spin coated on the glass coverslip and used for cell adhesion studies.
3. RESULTS AND DISCUSSION TE combines the principles of engineering with life science to develop biological substitute of extracellular matrix (ECM). 584
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Figure 2. (A) Design rationale for generating hydrophobic and hydrophilic surfaces. (B) Structures of Lys-appended diacetylene monomers with different amino protecting groups; and chemical structures of polydiacetylenes P1−P5.
rating amide linkages at the terminals of acetylene. Our approach is based on noncovalent interactions, wherein the monomers are self-assembled in the solid state for polymerization under UV light. This approach will ensure topochemical control and molecular level precision on the morphology of the synthesized polymers. This method will not only have high reproducibility but also have molecular level control on the morphology of the resulting scaffold. We have chosen lysine as an anchor unit due its diamine functionalities that can be masked or unmasked easily by using Fmoc and Boc protecting groups. The incorporation of diacetylene linkage through C-terminal of Lys provides a symmetrical structure endowed with similar or orthogonal protecting groups (Figure 2A). The advantage of this system is easy masking and unmasking of N-protecting groups to impart hydrophobic and hydrophilic properties to the surface. The diacetylene moiety allows the molecule to polymerize in the presence of UV light to obtain a polymer scaffold. Differently protected Lys was coupled with propargylamine to afford Lysderivatized with an alkyne moiety (Figures S1−S3 and S7−S9). Bis-Boc-L-Lys propargylamide was used to prepare the diacetylene D1 (Figures S4−S6), whereas Fmoc-Nε-Boc-L-Lys propargylamide was used to synthesize D2 (Figures S10−S12). Base labile Fmoc and acid labile Boc groups in D2 provide an opportunity to perform the selective deprotection of the Lys side chain without affecting the α-amino protecting group and vice versa. The diacetylenes D1 and D2 are good candidates for light-induced polymerization to yield PDAs P1−P5 (Figure 2B, Schemes S1 and S2).
The main aim of our work is to design and synthesize a synthetic equivalent of the ECM. The ECM is composed mainly of glycoprotein and is a highly cross-linked gel.17 It gives support and helps in navigation of extracellular signals. Typically, when cells are placed on a foreign surface outside human body, other than its ECM, it may survive with different characteristics or die. Only limited organs in the human body have the capacity to regenerate (e.g., liver and kidney) when damaged, that too if the ECM is intact. This led to intense efforts to develop synthetic ECM. The creation of tissues for replacing damaged tissues is likely to advance the medical field since TE allowed the fabrication of tissues such as skin, cartilage, ligament, tendon, bone, liver, and esophagus. However, an ideal scaffold fabrication is still not fully realized due to difficulties in fabricating materials with enough mechanical strength and interconnection channels. In order to mimic a natural bioenvironment, which resembles gel-like and cross-linked architecture of ECM, we designed a biomolecule-based polymeric system for TE. We envisioned that biomolecular synthons designed for directed self-assembly and further polymerization may provide nanostructure and cross-linked polymeric architecture suitable for TE. Typically materials with nanofibrous morphology can mimic the ECM ideal for tissue growth.18 Topochemical polymerization was considered by us as a method of choice, since it requires defined assembly by which the diacetylene monomers come in the close proximity of ∼5 Å before polymerization.19 The required registry for the topochemical polymerization is achieved by self-assembly of diacetylene monomers. Self-assembly is facilitated by incorpo585
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Figure 3. AFM images for (A) P1 and (B) P2; SEM images for (C) P3 and (D) P4.
P4, and P5 are insoluble, possibly due to the presence of high molecular weight polymers. With the monomers and polymers in hand, we investigated their self-assembly properties and the resulting surface morphologies by AFM, TEM, and SEM. D1 and D2 showed vesicular structure in ultramicroscopic analysis with a diameter 700−750 nm (Figures S20 and S21). The fibrous nature of the polymers was further confirmed by TEM and SEM (Figures 3, S22, S23, and S24). Microscopic analysis of polymers P1, P2, P3, and P4 showed long fibers with interstitial pores (Figures 3). Analysis of the fibers of P1 showed a “beads-on-chain” like architecture with a fiber thickness of 60−70 nm. The “beadson-chain” like morphology might have arisen from fusion of vesicles of D1 during polymerization. P2 fibers are thicker compared to P1, with a thickness in the range of 140−170 nm. P3 and P4 showed highly entangled fibers having a thickness in the nanometer dimension (Figures 3 and S24). An important feature of the polymer scaffolds that might help in cell adhesion is their wettability. Polymers P1 and P2 are hydrophobic due to the protecting groups, whereas P3, P4 and P5 are hydrophilic, having free amines along the polymer structure (Figure 2B). The wettability was analyzed by sessile drop water contact angle measurements (CA). A film of P1 showed a contact angle of 84° indicating a somewhat hydrophobic character. The Fmoc derivative P2 showed a CA of 136° consistent with a very hydrophobic surface. In contrast, films of deprotected derivatives P3, P4, and P5 with primary amino groups showed low CA, indicating hydrophilic surfaces (Figure S25 and Table 1). Mammalian cells do not have any specific receptors for polyamino acids, but cell membranes are endowed with negative charges, which can electrostatically engage polycationic polymers. Indeed, poly-Lys is known to support the adhesion of mammalian cells by means of electrostatic interactions between anionic sites on the plasma membrane components and cationic sites on the polymer surface.22 Films of the protected (P1 and P2) and deprotected polymers (P3, P4, and P5) were
Polymers P1 and P2 were prepared by irradiation of the corresponding diacetylenes D1 and D2 under a UV light from an Hg lamp for 3 min. In P1, all amino groups were protected by Boc groups, whereas in P2 the α-amino groups were protected by Fmoc and the ε-amino groups by Boc. In P3, αamino groups were protected by Fmoc and the ε-amino groups were free. P3 was prepared by the removal of Boc group from P2 using 25% trifluoroaceticacid (TFA) in dichloromethane. In P4, ε-amino group was Boc-protected and α-amino groups were unprotected. P4 was prepared by removal of the Fmoc group from P2 using 20% piperidine in dimethylformamide. Finally, P5 was prepared from P1 by the complete removal of Boc groups using TFA in dichloromethane. 1H NMR confirmed the formation of the polymers P1−P5 (Figures S13−S16). D1 underwent polymerization in the gel state and in the solid state under UV light to get P1. The color changed from colorless to pink color, which is indicative of the formation of polydiacetylenes (PDA). P1 was N-deprotected to get P5. D2 underwent polymerization under normal daylight, indicating a better packing in the solid state. The pink color of the PDA polymers indicated an ordered structure with a nonplanar geometry of the PDA chain.20 The attached groups on the diacetylene unit has a central role in controlling the planarity of the chain, and therefore the color of the polymer. The bulky and flexible side chain of Lys exert steric hindrance; therefore, the PDA chain adopts a nonplanar geometry. The formation of PDA was confirmed by Raman spectroscopic and gel permeation chromatographic (GPC) analysis. Raman spectra of polymers (Figures S17 and S18) showed a band at 2113 cm−1 (CC stretching) and 1509 cm−1 (CC stretching), indicative of the formation of PDA.21 The GPC was done on P2 since it is the only polymer that was soluble in THF. GPC analysis of the polymer P2 (Figure S19) showed a molecular weight of 40324 Da with a PDI of 1.02 and a degree of polymerization of 40. The other polymers P1, P3, 586
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proliferating only on polymers P1 and on glass coverslip; cell numbers increased after 72 h by a further 49% onto polymer P1 and 54% onto glass coverslip over the numbers after 48 h. In contrast, cell numbers increased by only 4% onto P2, 2% onto P3, and 2% onto P4. Thus, even if there were significantly more cells attached on polymers P3 and P4 after 4, 24, and 48 h compared to polymer P1, there was no difference between polymers P1, P3, and P4 after 72 h. Therefore, DPSC adhesion seems to be the most efficient on polymers P3 and P4, but cells have a limited proliferation rate on these polymers. We assessed cell spreading and cytoskeletal organization onto the four polymers P1−P4 after periods of 4, 24, 48, and 72 h (Figure 5). P5 was not studied since it did not form a stable film on the support surface. The organization of stress fibers (composed of F-actin bundles) was used for identification of cell shapes into round or branched and spindle morphology; round cells were defined considering circularity, while spindle cells were more elongated. Branched cells were characterized by more than two branches in the cell body.24 Cell spreading on the polymers P3 and P4 was evident after 24 h and comparable to glass coverslip (used as a reference for cell spreading). However, DPSC on P4 were more elongated than on P3 or on the glass coverslip and seemed to be oriented along the polymer fibers (in Figure 5D, note the overlay of fluorescence and bright-field microscopy, the latter revealing the polymer fibers). Most of the cells on P1 and P2 were round cells with poor cytoskeletal organization. After 72 h, most DPSC on P1 were branched with well-organized actin fibers. DPSC observed on P2 were either round or spindly with limited organized actin fibers. Among the various polymers, P3 was the most efficient for cell spreading, with branched cells and organized actin fibers, even though cells were not as well spread as on glass coverslip. DPSC on P4 were stretched and elongated, without branching, and with actin fibers, they were well-aligned along with polymer fiber orientation. Cell spreading can be described in terms of a balance between cellular forces of protrusion, contraction, and adhesion.25 We observed that cell spreading increased for polymers P1−P3 with increasing wettability after 72 h. Indeed, on P4, cells began aligning along single fibers instead of attaching to a wide area or multiple fibers. This effect had
Table 1. Sessile Drop Water Contact Angles of Films of Polymers P1−P5 polymer
contact angle (average)
surface groups
P1 P2 P3 P4a P5a
84.0 ± 0.14° 136.0 ± 1.33° 32.0 ± 3.60° 13.6 ± 5.7°
Nα-Boc, Nε-Boc Nα-Fmoc, Nε-Boc Nα-Fmoc, NH2 NH2, Nε-Boc NH2, NH2
a CA was difficult to measure reliably, as the drop quickly spread on the polymer surface.
hence probed in terms of cell attachment, morphology, and viability, given the potential of these materials as scaffolds for stem cell transfer. We assessed stem cell attachment onto the five polymers (P1−P5) after periods of 4, 24, 48, and 72 h. As a cell source, we used human dental pulp stem cells (DPSC) due to their strong potential mesenchymal stem cells in regenerative medicine.23 DPSC viability on different polymers was assessed via live/dead staining after 4 h in culture; fluorescein diacetate (FDA) was used to label the live cells (green) and propidium iodide (PI) to label the dead cells (red). The results show that all the attached cells were viable (stained in green) on all polymers, regardless of the wettability, confirming the absence of cytotoxicity (Figure 4A−E). The number of cells counted on each polymer, after fixation and nuclear staining, is presented in Figure 4H. We considered only polymers P1−P4. Indeed, the polymer P5 was not studied since it was washed away with the cell culture medium during the experiment. The polymer P5 is highly hydrophilic, and its nonadherence to the surface might be attributed to the presence of more abundant exposed amino groups compared to P3 and P4. There was no statistical difference between cell numbers on polymers P1 and P2, and between polymers P3 and P4, after 4 h and also after 24 h. Initial cell density (assessed after 4 h) on polymers P3 and P4 was higher compared to glass coverslip (p = 0.019 and 0.015, respectively). In contrast, polymers P1 and P2 showed lower cell numbers compared to glass coverslip (p = 0.026 and 0.023, respectively). Concerning cell proliferation after 48 h, DPSC kept
Figure 4. DPSC culture on polymers P1−P5. (A−G) Fluorescence microscopy after 4 h in culture followed by staining with FDA (green, vital staining) and PIES (red, dead cells). (A) polymer P1, (B) polymer P2, (C) polymer P3, (D) polymer P4, (E) polymer P5, (F) glass coverslip (negative control), (G) toxic control (DPSC rinsed with 4% formaldehyde). Scale bar = 100 μm. (H) Graph showing the cell counts of DPSC attached on polymers P1−P4, after 4, 24, 48, and 72 h of incubation (right side). Cells were counted per surface measuring 1400 × 1050 μm2. Experiments were made in triplicate, with five measurements per sample. Cell counts were plotted as mean ± standard error of the mean. 587
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Figure 5. Cytoskeletal organization of DPSC on polymers P1−P4, after 24 and 72 h of incubation (left and right pictures, respectively). Cells were stained with Hoechst (blue, nuclei staining) and phalloidin (red, actin cytoskeleton staining). (A) polymer P1, (B) polymer P2, (C) polymer P3, (D) polymer P4, (E) glass coverslip control. On images A−C, the blue/green background colors are due to autofluorescence of polymers. On image D, cells are overlays with bright-field to reveal the fibrous structure of P4. Magnification ×20 (scale bar = 90 μm).
Figure 6. Osteogenic differentiation assay: DPSC cultured on the various polymers in osteogenic medium for 3 and 6 days (left and right pictures, respectively). A glass coverslip was used as control. Cells were stained with Hoechst (blue, nuclei staining) and alkaline phosphatase antibodies (red, ALP staining). Background colors (green-yellow) are due to autofluorescence of polymers: (A) polymer P1, (B) polymer P2, (C) polymer P3, (D) polymer P4, (E) glass coverslip. Scale bar = 30 μm.
already been observed with electrospun PCL scaffolds when the space between fibers was 15−20 μm.26 To study DPSC differentiation on different polymers, cells were grown on polymers in osteogenic medium for six days. Alkaline phosphatase (ALP) activity was assessed by immunostaining with anti-ALP antibodies. ALP is an early differentiation marker expressed in differentiated cells producing mineralized matrix.27 After 3 days of osteoinduction, some fluorescence signal (ALP staining) was visible on polymer P4, but not on the other polymers. After 6 days, a clear staining was apparent on cells grown on polymer P4, while the staining on the control glass coverslip was faint (Figure 6). No mineralized matrix could be seen on either P1, P2, or P3, even after 6 days. These results indicate that cells on scaffold P4 are more differentiated at earlier time points than the cells grown on P1, P2, P3, and glass coverslip. The molecular architecture of
polymer P4 in terms of chemistry and topography appears to be osteoinductive for DPSC.28 In this study, we demonstrated that a diacetylene core with Lys at the termini is a versatile building block for molecular selfassembly and subsequent PDA polymer formation with tunable hydrophobic and hydrophilic surfaces. DPSC adhesion seems to be the most efficient on the hydrophilic polymers P3 and P4, compared to P1 and P2, but cells on the latter polymers proliferate faster. Comparing the hydrophobic polymers P1 and P2, P1 was found to be a better candidate than P2, judged by cell adhesion, spreading, and proliferation. This might be attributed to the porous and more fibrous nature of P1 compared to P2. Similarly, P4 is found to be a better scaffold than P3 for the same reason. Therefore, the molecular architecture plays a crucial role in cell attachment and spreading. The molecular architecture and the hydrophilicity 588
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of P4 appear to induce DPSC attachment and osteogenic differentiation. Therefore, P4 is a potential candidate as a selfassembled scaffold for stem cell therapy and TE. Poly(L-lysine) has poor mechanical properties and is therefore used in combination with other polymers for TE applications. The combination of lysine and PDA backbone increases mechanical strength and produces a more stable scaffold for TE. This method combines the advantages of cell adhesion properties of lysine and rigidity of synthetic PDA backbone. Further, the incorporation of peptides may improve the biodegradability of these PDA-based materials.
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ASSOCIATED CONTENT
* Supporting Information S
Detailed description about the synthetic procedures, 1H NMR, 13 C NMR, IR, HRMS, and microscopic analysis of the compounds. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected];
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We thank the Department of Science and Technology (DST and DST-FIST) New Delhi for financial support. S.S. thanks the Endeavour Awards Program (Australia) for an Endeavour Research Fellowship and CSIR (India) for the financial support.
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