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Preparation and characterization of a novel sodium alginate incorporated self-assembled Fmoc-FF composite hydrogel
Materials Science and Engineering C 58 (2016) 478–486
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Materials Science and Engineering C journal homepage: www.elsevier.com/locate/msec
Preparation and characterization of a novel sodium alginate incorporated self-assembled Fmoc-FF composite hydrogel Xiao Gong a,b, Christopher Branford-White c, Lei Tao a, Shubai Li d, Jing Quan a, Huali Nie a,b,⁎, Limin Zhu a,⁎⁎ a
College of Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, PR China State Key Laboratory for Modification of Chemical Fibers and Polymer Materials, Donghua University, Shanghai 201620, PR China Institute for Health Research and Policy, London Metropolitan University, London N78 DB, UK d Changzhou Institute of Engineering Technology, Changzhou 213164, PR China b c
a r t i c l e
i n f o
Article history: Received 21 April 2015 Received in revised form 8 August 2015 Accepted 27 August 2015 Available online 2 September 2015 Keywords: Self-assembly Peptide Sodium alginate Hydrogel Structure controllable
a b s t r a c t Dipeptides and their derivatives have attracted tremendous attention owning to their excellent abilities of selfassemble assembling into various structures which have great potentials for applications in biology and/or nanotechnology. In the present study, we dedicate to fabricate a rigid and structure controllable Fmoc-FF/SA composite hydrogel. We found that the modified dipeptide, fluorenyl-9-methoxycarbonyl (Fmoc)-diphenylalanine (Phe-Phe) can self-assemble into rigid hydrogels with structures of nanowires, layered thin films or honeycombs as the change of sodium alginate (SA) concentration. Meanwhile, CD-spectroscopy demonstrated that SA appeared to control the process, but it did not change the arrangement of the Fmoc-FF peptide. Our results demonstrated that the formed hydrogel showed physical and chemical stability as well as possessing good biocompatibility. Rheological measurements showed that the addition of SA could improve the stability of the hydrogel. Cell viability assay revealed that the Fmoc-FF and Fmoc-FF/SA hydrogels are both beneficial for cell proliferation in-vitro. Our results indicated that the fabricated Fmoc-FF/SA composite hydrogels could be used in tissue engineering and drug delivery in the future. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Recently, considerable improvements have been proceeded in the researches of molecular self-assembly systems over the past decades, as is reported, the systems include macromolecules [1,2], proteins [3], nucleic acids [4,5], peptides [6,7], liposomes [8] and graphite oxides [9, 10]. At the same time, peptide-based systems being capable of selfassembling, environment friendly, engaged in molecular recognition and diverse functionality, and have enormous application potential in biological and non-biological fields [11–15]. These molecules usually form hierarchical structures at both nano- and micro-scales. It is assumed that the assembly ordered of these building blocks into defined nanostructures was relied on specific molecular recognition sites. These are facilitated by strong covalent bonds or a combination of week non-covalent interactions, including hydrogen bonds, Vander Waals forces, hydrophobic forces, electro-static interactions, π–π stacking, and chiral dipole–dipole interactions [16,17].
⁎ Correspondence to: H. Nie, College of Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, PR China. ⁎⁎ Corresponding author. E-mail addresses: [email protected] (H. Nie), [email protected] (L. Zhu).
http://dx.doi.org/10.1016/j.msec.2015.08.059 0928-4931/© 2015 Elsevier B.V. All rights reserved.
Peptide self-assembly was first certified by the characterization of the repetitive 16-residue peptide motif n-AEAEAKAKAEAEAKAK-c (EAK 16-II), a fragment of a left-handed Z-DNA binding protein in yeast [18]. After that, more peptide-based materials capable of selfassembling were discovered or synthesized including amphiphilic peptides [19,20], cyclic peptides [21–23], dendritic peptides [6], aromatic dipeptides [24], surfactant-like oligo-peptide [25,26], copolypeptides [27] and the like. Moreover, a class of peptide building blocks sharing common features (RDA16-I, RAD16-II, EAK-I) were designed, and they all showed great potential in facilitating 3D tissue cultures mainly due to the formation of self-assembled nanofiber scaffolds [15]. Among these peptide-based materials, diphenylalanine (FF) and its derivatives have often been used as a model structure for evaluating molecular mechanisms of self-assembly systems. Morphological observations and other approaches have been utilized to investigate the effect of external stimuli on FF self-assembly. These investigations include the effect of role of solvent, assembly surface and pH [28,29]. It was proved that these colloids were not only stimuli-responsive to pH and temperature, but also showed novel and adaptive encapsulation properties [30]. For example, FF-based building blocks are capable of assembling into various functional nano-structures, such as nanotubes [31–34], spherical vesicles [35], nanofibrils and nanoribbons [19,36], nanowires [37,38] and ordered molecular chains [39]. Moreover, peptide hydrogels and their assembly processes have been considered as good candidates
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in biological applications, involving tissue engineering [40–44], drug delivery [45–47], bio-imaging and biosensors [48–50]. Although there have been lots of studies on peptide-based hydrogels, we still knew little about the biological mechanisms of Fmoc-peptide hydrogels, and there were still many shortcomings of the hydrogels in biological safety, cost savings, strength, range of applications and so on. For example, Huang et al. introduced konjac glucomannan (KGM) into the Fmoc-FF system to prepare a potential carrier for drug delivery, but it took nearly 200 min to form a rigid transparent hydrogel [51]. For another example, the frozen glucono-δ-lactone (Gdl) in Fmoc-FF system indeed resulted in the formation of macro-porous hydrogels, however, the cryogels turned mechanically weaker (G′ b 1000 Pa) than the Fmoc-FF hydrogels prepared under the same condition [52]. As a natural polysaccharide, SA is nontoxic, with good solubility and fast gelation propriety, and has been widely applied to the field of food, medicine, etc. Shi et al. prepared a novel and superporous SA-based hydrogel by the self-assembling micelle templating [53]. And KGM/SA hydrogels were prepared using graphene oxide as a drug-binding effector for cancer therapy drug loading and release [54]. In previous studies, scientists introduced polysaccharides into the system to regulate peptide-based hydrogels either from intensity or the structure, but few have shown the capacity to both ensure the strength and the structure of the hydrogels in a short time. The aim of this study is to find a novel and efficient approach to prepare rigid and structure controllable Fmoc-FF/SA composite hydrogels with stable physical and chemical properties. It is hopeful that our study will contribute to understanding the mechanism of peptide selfassembly and the role of polysaccharide in the peptide selfassembly process.
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2. Experimental 2.1. Materials and methods The Fmoc-diphenylalanine (Fmoc-Phe-Phe-OH, Fmoc-FF) peptide in lyophilized form was purchased from Bachem (Bubendorf, Switzerland). Sodium alginate (AR, MW: 398.31) was obtained from Aladdin Chemistry Co. Ltd. Dimethyl sulfoxide (DMSO) was purchased from the Sinopharm Chemical Reagent Co. Ltd. (Shanghai, China). Porcine pepsin (250 units mg−1) and trypsin (3194 units mg−1) were purchased from J&K Scientific Ltd. (Beijing, China). Dulbecco's Modified Eagle Medium (DMEM) was purchased from Genome Biological Technology Co. Ltd. (Hangzhou, China). Penicillin and streptomycin were purchased from Generay Biological Engineering Co. Ltd. (Shanghai, China) and fetal calf serum was purchased from Life Technologies Corporation (America). Other reagents were obtained from the Beijing Chemical Reagent Corporation and were used as received. Water was distilled in a three-stage Millipore Milli-Q plus 185 purification system and had a resistivity N18.2 MΩ. 2.2. Hydrogel preparation SA powders were gradually dispersed in double deionized (dd) H2O (1.0–10.0 mg ml− 1) while stirring, and then stirred for 12 h at room temperature to fully dissolve them. Fmoc-FF stock solution was freshly prepared by dissolving the peptides in DMSO (100.0 mg ml−1) before use. The peptide stock solution was then diluted to a final concentration of 2.0 mg ml−1 by adding dropwise to ddH2O and SA solutions, respectively. The mixture solutions were left at room temperature for three
Fig. 1. A schematic illustration of the gelation process and stability tests of the hydrogel. (a) Molecular structure of Fmoc-FF; (b) white floc emerged after adding peptides; (c) white turbid viscous liquid after sufficient shake; (d) formation of the translucent hydrogel; (e) macroscopic image of the hydrogel samples after ultrasonic processing for 2 h; (f) macroscopic image of the hydrogels after shaking in the shaking bath at 37 °C with 1.0 ml PBS solution; (g) the remained quantity of Fmoc-FF peptide after in vitro digestion of Fmoc-FF hydrogel in simulated gastric fluid; (h) the remained quantity of Fmoc-FF peptide after in vitro digestion of Fmoc-FF hydrogel in simulated intestinal fluid.
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days after sufficient mixing on a miniature oscillator, leading to the formation of the highly transparent Fmoc-FF peptide and Fmoc-FF/SA hybrid hydrogels [6,51]. In order to distinguish the different samples, the Fmoc-FF hydrogel and Fmoc-FF/SA composite hydrogels were labeled 1# (SA, 0 mg ml−1), 2# (SA, 1.0 mg ml−1), 3# (SA, 3.0 mg ml−1), 4# (SA, 5.0 mg ml−1), 5# (SA, 7.0 mg ml−1), and 6# (SA, 10.0 mg ml−1), respectively. 2.3. Hydrogel stability To evaluate the stability of the hydrogels, 0.2 M PBS, pH 7.4 (2.0 ml), containing 0.1% (w/v) NaN3, were added to the upside of isopyknic Fmoc-FF and Fmoc-FF/SA (both 2.0 mg ml−1) hydrogels, respectively. Samples were placed in a rotary shaker at 70 rpm and 37 °C for 12 h and morphological changes were recorded with a digital camera (COOLPIX S80, Nikon Corporation, Japan). 2.4. In vitro digestion The stable hydrogels were immersed in simulated gastric (0.5 ml, 10 mM PBS–HCl, pH 2.0, 20 μg ml−1 pepsin) and simulated intestinal fluids (0.5 ml 10.0 mM PBS, pH 7.4, 20 μg ml−1 trypsin), respectively [51]. The hydrogels were then placed in a rotary shaker, 150 rpm and 37 °C for 24 h. Enzymatic reactions were deactivated by placing samples in boiling water bath for 15 min after centrifugation at 15,000 rpm for 5 min. The supernatants were collected and the amount of Lphenylalanine was determined using the high-performance liquid chromatography (HPLC, Waters, 2489, Waters Corporation, USA) and Agilent TC-C18 column. HPLC was performed at 210 nm using a mobile phase (water: acetonitrile (70:30, v/v) at a flow rate of 1.0 ml min−1, and 20 μl samples were injected. The amount of Fmoc-FF peptide
remained was calculated by removing the amount of L-phenylalanine determined.. 2.5. Rheological measurement In-situ hydrogel formation and rheological properties were measured using an ARES-RFS high rotational rheometer (TA Instruments, USA) at 25 °C with a paralleled plate of geometry (50 mm in diameter, tool inertia 62.5 g·cm2, max auto-tension displacement 3.0 mm, minimum applied dynamic force = 1.0 gmf). Time sweep oscillatory measurements were performed immediately after shaking the samples and preparing mixtures (30 s), diluting the Fmoc-FF stock solutions to give a final concentration of 4.0 mg ml− 1 in ddH2O and SA (1.0 mm gap, fluid density 1.0 g cm−3, strain 0.5%). Whereafter, dyn strain frequency sweep (0.1–100 Hz, strain 0.5%) measurement was conducted to determine the frequency dependent storage and loss sheer moduli, where in the time sweep oscillatory test was performed [51,52]. All tests were repeated three times and the average value was taken. 2.6. Scanning electron microscopy (SEM) To gain insight into the real morphology of the self-assembled structures of the (composite) hydrogels, all the samples (3 days old) were prepared by rapid freezing for 2 h at − 80 °C, and drying for 24 h in a freeze drier (Christ Alpha1-2, Osterode, Germany). In order to observe the self-assembled morphology of Fmoc-FF in the composite hydrogels, the samples were dispersed in water and SA was removed by membrane filtration. All the samples were sputter-coated and then images were recorded using a JSM-5600LV scanning electron microscope (JEOL, Tokyo, Japan).
Fig. 2. Dynamic rheology analysis of self-assembled Fmoc-FF hydrogel (1#) and Fmoc-FF/SA composite hydrogel (2#). (a) Time dependence of G′ and G″ of Fmoc-FF peptide hydrogel; (b) time dependence of G′ and G″ of Fmoc-FF/SA composite hydrogel; (c) the change in G′ and G″ of Fmoc-FF peptide hydrogel with the increased frequency (0.1–100 Hz); (d) the change in G′ and G″ of Fmoc-FF/SA composite hydrogel with the increased frequency (0.1–100 Hz).
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2.7. Transmission electron microscopy (TEM)
2.10. Circular dichroism spectroscopy (CD)
After freezing–drying, Fmoc-FF and Fmoc-FF/SA xerogels were negatively stained with phosphotungstic acid (1%) for 2 min. Then a drop of the sample xerogel was placed on a 400-mesh copper grid and dried at room temperature. Samples were viewed at 80 kV using a JEM-2100 electron microscope (JEOL Ltd., Japan).
CD spectra were recorded using a Jasco J-810 CD spectrometer (Jasco, Glasgow, UK), 190–260 nm, with a resolution of 0.5 nm at room temperature. Samples were prepared by dispersing the xerogels in aqueous solutions to avoid DMSO interfacing. The path length of the quartz cell was 1.0 mm and 0.2 mg ml− 1 [51,52]. All tests were repeated three times and the average value was taken.
2.8. Thermogravimetric analysis (TGA) 2.11. Viability analysis TGA (TA 2050, America) was conducted after the freezing–drying process. The samples were heated at the rate of 8 °C min− 1 under nitrogen. 2.9. Fourier transform infrared spectroscopy (FTIR) FTIR spectra of Fmoc-FF/SA xerogels, Fmoc-FF and SA were undertaken on a Nicolet-6700 FTIR spectrometer (Thermo Fisher, USA) over 3750–600 cm−1 at a resolution of 2 cm−1; 32 scans were taken.
Studies on the scaffold effects of the composite hydrogels for tissueengineering applications and in-vitro cellular experiments were undertaken using the following protocol. Fresh Fmoc-FF and Fmoc-FF/SA composite hydrogels (300 μl) were prepared in water and SA, respectively. After autoclaving, samples were placed in a 24- hole culture plate system (Corning) and then fumigated with alcohol overnight. For cell-growth pig iliac endothelial cells (PIEC) were grown in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10%
Fig. 3. SEM images of self-assembled Fmoc-FF peptide hydrogel and Fmoc-FF/SA composite hydrogels. (a) Fmoc-FF peptide hydrogel; (b)–(f) Fmoc-FF/SA composite hydrogels with different concentrations of sodium alginate (1.0, 3.0, 5.0, 7.0, 10.0 mg ml−1, respectively).
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fetal calf serum, 100 U ml−1 penicillin and 100 U ml−1 streptomycin. Cells were maintained at 37 °C in a humidified atmosphere containing 5% CO2. Subconfluent cells were harvested by trypsinization, counted, and diluted in the cells media to 1.0 × 106 cells ml−1. Before cell implantation, the hydrogels were washed (three times) with PBS (pH 7.4, 0.01 M) and DMEM, respectively. Then 20 μl of PIEC (2 × 104 cells) was over laid on the hydrogels. After 24 h incubation at 37 °C with 5% CO2 in the constant temperature incubator, the viability of the cells was determined using an MTT assay [6,55]. The cell medium was replaced daily and the results from triplicate samples taken. To qualitatively confirm cell viability and proliferation upside the hydrogels, confocal laser scanning microscopy (CLSM, Carl Zeiss LSM 700, Germany) was used. Cover slips with a diameter of 14 mm were pretreated with 5% HCl, 30% HNO3, and 75% alcohol and then fixed in 24-well tissue culture plate. Total number of 2 × 104 PIEC was seeded per well upside the hydrogels and cultivated for 7 days. Then the cells were rinsed with PBS for 3 times fixed with glutaraldehyde (2.5%) for
15 min at 4 °C, and stained with rhodamine (1 μg ml−1) for 15 min at 37 °C using a standard procedure. Finally, all samples were washed with PBS and observed via CLSM. 3. Results and discussion 3.1. Hydrogel preparation The Fmoc-FF hydrogel was prepared according to the following method: First, dissolve the Fmoc-FF peptide in DMSO (Fig. 1a), then inject it into ddH2O (Fig. 1b). Meanwhile, the Fmoc-FF turned into white floc immediately (Fig. 1c). Whereafter, shake the sample tube to mix evenly. Within a few minutes, a stable transparent hydrogel formed (Fig. 1d) from a state of white turbid colloid. The Fmoc-FF/SA composite hydrogel was prepared by mixing the peptide with SA at different concentrations. The gelation of the Fmoc-FF was also accompanied by the formation of a transparent hydrogel, though it took a little longer time
Fig. 4. TEM and SEM micrographs of self-assembled Fmoc-FF peptide hydrogel and Fmoc-FF/SA hydrogels. (a) TEM micrograph of Fmoc-FF; (b)–(d) TEM micrographs of Fmoc-FF/SA composite hydrogels (3#, 5#, 6#); (e) SEM image of stacked massive structure of Fmoc-FF peptide; (f) SEM image of membranous Fmoc-FF structure after removal of SA (5#).
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3.3. Rheology characterization
Fig. 5. TG curves of (a) the dried sodium alginate; (b) Fmoc-FF/SA composite hydrogel (2#); and (c) Fmoc-FF.
(ranging from several minutes to several quarters) to form a stable and transparent composite hydrogel as the concentration of sodium alginate increased.
3.2. Stability tests The stability of the hydrogels was assessed by ultrasonic concussing and incubation at pH 7.4 and 37 °C. The change of the two hydrogel samples was noted after they had been treated by ultrasonication for 2 h. It was clear that both the Fmoc-FF hydrogel (1#) and Fmoc-FF/SA composite hydrogels (2#–6#) remained almost unchanged (Fig. 1e). As the concentration of SA increased, the state of hydrogel systems gradually changed from turbid viscous fluid to samples with clear “water layer–gel layer” interface (Fig. 1f), showing preferable properties of anti-phosphate buffer. These results also illustrated that the interference of SA enhanced the stability of the hydrogel system. In the presence of proteases (pepsin and trypsin), in-vitro digestion of the FmocFF peptide and the composite hydrogels were evaluated. No detection of phenylalanine was found after 24 h and it happened because these structures were resistant to proteolytic attack (Fig. 1g and Fig. 1h). The results indicated that this system could probably provide as a platform for future applications of oral drugs and their delivery [6].
The mechanical properties of the hydrogels are important for assessing their suitability for future applications. Thus dynamic rheology experiment was undertaken to investigate the kinetic formation of hydrogel and the effect of SA on the gelation process. Fig. 2a showed the storage modulus (G′) and shear modulus (G″) change over time. G ′ and G″ were almost invariable within 60 s. However, from 60 s to 150 s, the values both rose sharply and reached a plateau at about 150 s. From the date, the storage modulus G′ (~ 4300 Pa) exceeded that of the loss modulus G″ (~400 Pa) by ~11 times, indicating the formation of a steady viscoelastic hydrogel [37,51]. However, the gelation time (i.e. the time after which G′ becomes larger than G″) of Fmoc-FF/ SA (2#) was much shorter than the Fmoc-FF hydrogel and was even difficult to record (Fig. 2b), though the stable Fmoc-FF/SA composite hydrogel formed only with a G′ of 2000 Pa, and G″ of 200 Pa. This result was also consistent with the findings that the rigid transparent FmocFF hydrogel formed within a few minutes, but a longer time for the Fmoc-FF/SA composite hydrogel. It is assumed that the steric hindrance of SA, and strong hydrogen bonds between the molecules, impeded the movement of Fmoc-FF molecules and their aggregation, thereby, prolonged the process of self-assembly. Nondestructive frequency sweep indicated that the shear modulus G ″ of Fmoc-FF dramatically decreased with frequency, especially when the frequency increased to 30 Hz, despite a slight increase of G′ for Fmoc-FF hydrogel between 0.1 Hz to 100 Hz (Fig. 2c). However, in contrast with Fmoc-FF hydrogel, both G′ and G″ of the Fmoc-FF/SA composite hydrogel were not affected by the increase of shear frequency (Fig. 2d). In short, the composite hydrogel (SA, 1.0 mg ml−1) exhibited better frequency stability and toughness than the Fmoc-FF hydrogel within the frequency range. That is because SA contains a large number of hydroxyl groups, making it easy to form intermolecular hydrogen bond. Accordingly, the enhanced hydrogen bonding among selfassembled peptide, water and SA (–OH groups) strengthen the stability of composite hydrogels.
3.4. Morphology characterization The morphological structures of the self-assembled hydrogels were observed by SEM and TEM, respectively. As shown in Fig. 3a, when the Fmoc-FF stock solution was directly injected into ddH2O, selfassembled blocky structure in micro-scale was formed, which differed from previous reports [6,24]. However, when SA (1.0 mg ml−1) was introduced into the Fmoc-FF system, the morphology of the assembled
Fig. 6. (a) FTIR spectra of Fmoc-FF hydrogel, sodium alginate and Fmoc-FF/SA composite hydrogel (2#); (b) CD spectra of Fmoc-FF peptide and Fmoc-FF/SA composite hydrogel (2#).
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structure, it was comprehensible that the Fmoc-FF hydrogel was the toughest among the all of hydrogels. The Fmoc-FF hydrogel finally self-assembled into a micro-sized blocky structure, while the Fmoc-FF/ SA composite hydrogels formed nanowire networks or “honeycomblike” structure etc. The presence of numerous cavities, perhaps contributed to the decrease of G′. TEM showed that Fmoc-FF hydrogel consisted of plenty of blocky structures (Fig. 4a, c) in similar morphology. Fmoc-FF in the composite hydrogel (3#) can self-assemble into long nanowires which are several microns in length and several tens of nanometers in width (Fig. 4b). The “honeycomb sheet” structure of the composite hydrogel (6#) was also confirmed by TEM (Fig. 4d). The reason why we did not observe a complete three-dimensional cellular structure was that the material was redissolved prior to TEM observation. Compared with the membranous structure of the composite hydrogel (5#, Fig. 4c), the “honeycomb sheet” was thinner and flexible with clear wrinkles on the surface. All these morphological observations illustrated that the SA played more important roles as an inducer or a template to immobilize Fmoc-FF.. Fig. 7. Cell viability tests on the surface of Fmoc-FF hydrogel and Fmoc-FF/SA composite hydrogels for 1, 3, 5, and 7 days.
hydrogel changed from blocky structure to short nanowires, tens of nanometers to several microns in length (Fig. 3b). Moreover, when we increased the concentration of SA to 3.0 mg ml−1, nanowires from several to tens of microns in length appeared (Fig. 3c). To further increase the concentration of SA (5.0 mg ml− 1), the nanowire structure was hardly observed and films containing a large number of folds generated (Fig. 3d). As shown in Fig. 3e, when SA of 7.0 mg ml−1 was applied, pieces of thin films constituted the entire hydrogel scaffold. Fig. 3f revealed that in the presence of high concentration of SA (10.0 mg ml−1), the composite hydrogels self-assembled into “honeycomb-like” structure that contained several cavities. It was reasonable to assume that the removal of water held within the honeycomb scaffold contributed to the structural stability and so shrank the “honeycomb sheet” to form a “shriveled and complicated” pile. Various structures were formed by controlling the concentration of SA, and this could be due to the steric hindrance of SA and the impact of strong non-covalent bond formation that facilitated Fmoc-FF assembly after aggregation, such as the strong hydrophobic interaction of the polypeptide, π–π stacking of the Fmoc-FF molecules and hydrogen-bond interaction with SA molecule [25,28]. From the perspective of hydrogel
3.5. Supramolecular arrangement TGA was performed to investigate the combination of Fmoc-FF and SA. As shown in Fig. 5a, the thermal decomposition of sodium alginate was divided into three stages (named Stages 1, 2, 3). Stage 1 (within 100 °C) was the effect of lactonization and transglycosylation; Stage 2 (from 220 to 250 °C) was fracture of the sodium alginate skeleton, and the final stage was the carbonization effect. For Fmoc-FF, only one stage of weight loss (from 220 to 300 °C) occurred, and this accounted for 90% weight loss (Fig. 5c). It was clear in Fig. 5b that the weight loss of Fmoc-FF/SA from 200 to 400 °C compromised that for sample SA and Fmoc-FF, indicating that the composite hydrogel sample integrated the properties of the two materials. FTIR was used to verify the above assumption. The spectrum of the Fmoc-FF/SA xerogel after freezing–drying (Fig. 6a) showed the position of the amide I band, a weak peak at 1692 cm−1 and a stronger one at 1652 cm−1, indicating the predominant β-sheet character [19]. It was clear that Fmoc-FF/SA composite hydrogel showed typical amide II and amide III adsorption peak centered at 1538 cm−1 and 1258 cm−1 that attributed to Fmoc-FF and carboxylate adsorption peak at 1606 cm−1 and 1414 cm−1 that attributed to SA. The presence of all these characteristic peaks confirmed a good blend of SA and Fmoc-FF,
Fig. 8. Confocal microscopic images of (a1–a3) Rhodamine stained PIEC upside of Fmoc-FF hydrogel; (b1–b3) Rhodamine stained PIEC upside of Fmoc-FF/SA composite hydrogel (2#).
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which laid a foundation for regulating the structure of self-assembled peptide without changing the arrangement of the Fmoc-FF. The secondary molecular structure of Fmoc-FF peptide was assessed using circular dichroism (CD). The spectra of Fmoc-FF peptide (Fig. 6b) were similar to that of the composite hydrogel, with positive peaks at ~ 194 nm, and ~ 214 nm, indicating the β-sheet arrangement of the Fmoc-FF molecular structure [24,34]. Random coil conformation of the Fmoc-FF molecular structure could be confirmed by a strong negative peak at 197 nm and a weak positive wide peaks at 217–223 nm. For the Fmoc-FF/SA composite hydrogel, the strong peak at ~198 nm, and the negative peak at ~215 nm demonstrated the β-sheets conformation, while the negative peak at 201 nm and the positive peak ~219 nm were due to random coiling [56]. Meanwhile, morphology of Fmoc-FF in Fig. 6b was also a good explanation of the role SA played: SA served as an inducer to assemble the composite hydrogel and a template to immobilize the Fmoc-FF. Previous studies reported that molecules with hydroxyl, benzene and other functional groups were capable of interacting with peptide molecules to form different self-assembled materials with various shapes and functions [48,57]. It is well known that SA owns a large numbers of hydroxyl groups, which would certainly influence the arrangement of the Fmoc-FF peptide due to the enhanced hydrogen bond among SA, water, and peptide. To study the effect of SA on the self-assembly of Fmoc-FF molecular, SA was removed from the xerogels by dissolving in ddH2O and vacuum filtration. It was confirmed that Fmoc-FF and SA have been mixed well together. If there is some interaction between Fmoc-FF and SA, the structure of the Fmoc-FF/SA composite hydrogel should be seriously damaged with the dissolution of SA. If SA just played a role of regulating agent, the self-assembled structures might be well preserved. However, compared with those without being washed, the morphology of selfassembled Fmoc-FF remained the same after the removal of SA (Fig. 4e, f). This phenomenon directly indicated that SA played an important role in “inducing agent” during the Fmoc-FF self-assembly process and didn't change the arrangement of the Fmoc-FF peptide [51]. Based on the above research, we can summarize that the role of SA in the Fmoc-FF/SA system mainly involves: (i) serving as an inducer to form the composite hydrogel and a template to immobilize Fmoc-FF; (ii) contributing to Fmoc-FF aggregation in situ and growing into desired structures through the steric hindrance of SA under the different concentration; (iii) stabilizing the 3D gel network structure due to the enhanced hydrogen bonding among self-assembled peptide, water and SA (–OH groups). 3.6. Cell viability assay To assess and evaluate the ability of composite hydrogels to be used in biological applications, cell viability was analyzed following 1, 3, 5, and 7 days of incubation period (Fig. 7). After one day, cells were taken from all peptide hydrogel surfaces, and they all increased noticeably, reaching nearly 50% growth rate compared with the control. After 3 days, the number of cells on the composite hydrogels with SA (1.0 mg ml−1–10.0 mg ml−1) was nearly identical to that Fmoc-FF hydrogel without SA (1#), which increased by ~ 77% compared with the control experiment, showing good cell viability. Over the next few days, cells continued to grow. Particularly, cells on the Fmoc-FF peptide hydrogel proliferated by 93% after 7 days of cultivation compared with the same stage of control. However there was a small decrease in growth rate of cells on the composite hydrogels after days 5–7, with the cell population increase of 70% compared with the control group. Therefore, viability assay showed that sodium alginate had little effect on cell growth. The survival status of PIEC cultured on the hydrogels was qualitatively confirmed via confocal microscopic imaging of the red fluorescence of rhodamine. As shown in Fig. 8a1–a3, when the Fmoc-FF hydrogel was inoculated with PIEC, cells grew rapidly and were fairly homogenous on the surface. Fluorescent photomicrographs also
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showed good morphology of PIEC in the composite hydrogel (Fig. 8b1–b3). By comparing Fig. 8b2 with Fig. 8b1 and b3, it was clear that cells grew not only on the surface of the composite hydrogel, but also in the internal space of the nanowire networks as the red arrows pointed to. It was also consistent with Fig. 7. In short, Fmoc-FF finally self-assembled into blocky structures in micro-scale, while the FmocFF/SA composite hydrogels formed nanowire networks or “honeycomb-like” structure, leading to the weaker of G′. However, from the perspective of applications, especially in tissue engineering as well as cell and/or tissue regeneration, Fmoc-FF/SA hydrogels with rich cavity structure could provide the cells adequate growth space. 4. Conclusions In this study, we designed a novel composite hydrogel by inducing SA into Fmoc-FF peptide system. The hydrogel was stable in buffer and proteolytic digestion. Rheological results revealed that the asprepared hydrogels were consistently intact and the morphological observations indicated that the Fmoc-FF/SA can self-assemble into nanowires, layered thin films, or honeycomb-like structures, while the Fmoc-FF peptide self-assembled into blocky structure. The driving force for the self-assembled nanostructure was inferred to be due to hydrogen bonding and aromatic packing factors. SA also played an important role in regulating the movement and combination of Fmoc-FF molecules during assembly. Data from CD implied that the hydrogels had similarities in terms of β-sheet and random coil peptides. Finally, cell viability assay demonstrated that Fmoc-FF/SA composite hydrogels exhibited good biocompatibilities and enhanced the growth of epithelial cells. It is expected that the novel composite hydrogels would be utilized in various broad applications in future tissue engineering as well as cell and/or tissue regeneration. Acknowledgments The project was funded by the Shanghai Natural Science Foundation (14ZR1401300), National Key Fund (EG2014021), Qing Lan Project, Changzhou Applied Science Foundation (CJ20120014) and State Key Laboratory for Modification of Chemical Fibers and Polymer Materials Donghua University. References [1] B.M. Blunden, H.X. Lu, M.H. Stenzel, Biomacromolecules 14 (2013) 4177–4188. [2] W.I. Hung, C.H. Chang, Y.H. Chang, P.S. Wu, C.B. Hung, K.C. Chang, M.C. Lai, S.C. Hsu, Y. Wei, X.R. Jia, J.M. Yeh, Langmuir 29 (2013) 12075–12083. [3] Y.P. Chuan, Y.Y. Fan, L.H.L. Lua, A.P.J. Middelberg, J. R. Soc. Interface 7 (2010) 409–421. [4] P.W.K. Rothemund, A. Ekani-Nkodo, N. Papadakis, A. Kumar, J. Am.Chem.Soc. 126 (2004) 16344–16352. [5] A. Gangar, A. Fegan, S.C. Kumarapperuma, P. Huynh, A. Benyumov, C.R. Wagner, Mol. Pharm. 10 (2013) 3514–3518. [6] A. Mahler, M. Reches, M. Rechter, S. Cohen, E. Rigid Gazit, Adv, Mater 18 (2006) 1365–1370. [7] N. Nakatsuka, S.N. Barnaby, K.R. Fath, I.A. Banerjee, J Biomat Sci-Polym E. 23 (2012) 1843–1862. [8] D.G. Yu, C. Branford-White, G.R. Williams, S.W.A. Bligh, K. White, L.M. Zhu, N.P. Chatterton, Soft Matter 18 (2006) 1365–1370. [9] C.M. Chen, Q.H. Yang, Y.G. Yang, W. Lv, Y.F. Wen, P.X. Hou, M.Z. Wang, H.M. Cheng, Adv. Mater. 21 (2009) 3007–3011. [10] C. Wu, X.Y. Huang, G.L. Wang, L.B. Lv, G. Chen, G.Y. Li, P.K. Jiang, Adv. Funct. Mater. 23 (2013) 506–513. [11] L. Liu, X. Liu, H. Deng, Z. Wu, J. Zhang, B. Cen, Q. Xu, A. Ji, J Biomat Sci-Polym E. 25 (2014) 1331–1345. [12] D. Silva, A. Natalello, B. Sanii, R. Vasita, G. Saracino, R.N. Zuckermann, S.M. Doglia, F. Gelain, Nanoscale 23 (2013) 506–513. [13] P. Kongsuphol, S.K. Arya, C.C. Wong, L.J. Polla, M.K. Park, Biosens. Bioelectron. 55 (2014) 26–31. [14] P. Moitra, K. Kumar, P. Kondaiah, S. Bhattacharya, Angew. Chem. Int. Ed. 53 (2014) 1113. [15] S.G. Zhang, Nat. Biotechnol. 53 (2014) 1113–1117. [16] J.E. Huheey, E.A. Keiter, R.L. Keiter, Inorganic Chemistry: Principles of Structure and Reactivity, Prentice Hall, Upper Saddle River, NJ, 1997. [17] P.Y. Bruice, Organic Chemistry, Prentice Hall, Upper Saddle River, NJ, 2003.
486
X. Gong et al. / Materials Science and Engineering C 58 (2016) 478–486
[18] S.G. Zhang, T. Holmes, C. Lockshin, A. Rich, P. Natl. Acad. Sci. USA 90 (1993) 3334–3338. [19] Y.Y. Lin, Y. Qiao, P.F. Tang, Z.B. Li, J.B. Huang, Soft Matter 7 (2011) 2762–2769. [20] J.D. Hartgerink, E. Beniash, S.I. Stupp, Science 294 (2001) 1684–1688. [21] M.R. Ghadiri, J.R. Granja, R.A. Milligan, D.E. Mcree, N. Khazanovich, Nature 366 (1993) 324–327. [22] L. Ruiz, S. Keten, J. Eng. Mech. 140 (2014) 431–442. [23] Y.Z. Wang, S.J. Yi, L.M. Sun, Y.J. Huang, S.C. Lenaghan, M.J. Zhang, J. Biomed. Nanotechnol. 10 (2014) 445–454. [24] M. Hughes, P.W.J.M. Frederix, J. Raeburn, L.S. Birchall, J. Sadownik, F.C. Coomer, I.H. Lin, E.J. Cussen, N.T. Hunt, T. Tuttle, S.J. Webb, D.J. Adams, R.V. Ulijn, Soft Matter 8 (2012) 5595–5602. [25] X.J. Zhao, S.G. Zhang, Chem. Soc. Rev. 35 (2006) 1105–1110. [26] Y.Y. Lin, T. Qiao, P.F. Tang, Z.B. Li, J.B. Huang, Soft Matter 7 (2011) 2762–2769. [27] T.J. Sanborn, P.B. Messersmith, A.E. Barron, Biomaterial. 23 (2002) 2703–2710. [28] X.H. Yan, Y. Cui, Q. He, K.W. Wang, J.B. Li, Chem. Mater. 20 (2008) 1522–1526. [29] R.L. Huang, W. Qi, R. Su, Soft Matter 7 (2011) 6418–6421. [30] X.H. Yan, P.L. Zhu, J.B. Fei, J.B. Li, Adv. Mater. 22 (2010) 1283–1287. [31] M. Reches, E. Gazit, Nat. Nanotechnol. 1 (2006) 195–200. [32] R.J.A. Hill, V.L. Sedman, S. Allen, P.M. Williams, M. Paoli, L. Adler-Abramovich, E. Gazit, L. Eaves, S.J.B. Tendler, Adv. Mater. 19 (2007) 4474–4479. [33] X.H. Yan, Y. Cui, Q. He, K.W. Wang, J.B. Li, W.H. Mu, B.L. Wang, Z.C. Qu-yang, ChemEur J 14 (2008) 5974–5980. [34] P. Bharagava, Y.F. Tu, J.X. Zheng, H.M. Xiong, R.P. Quirk, S.Z.D. Cheng, J. Am. Chem. Soc. 129 (2007) 1113–1121. [35] M. Reches, E. Gazit, Nano Lett. 4 (2004) 581–585. [36] I. Cherny, E. Gazit, Angew. Chem. Int. Edit. 47 (2008) 4062–4069. [37] T.H. Han, J. Kim, J.S. Park, C.B. Park, H. Ihee, S.O. Kim, Adv. Mater. 19 (2007) 3924–3927. [38] J. Ryu, C.B. Park, Adv. Mater. 20 (2008) 3754–3758. [39] M. Lingenfelder, G. Tomba, G. Costantini, L.C. Ciacchi, A. De Vita, K. Kern, Angew. Chem. Int. Ed. 46 (2007) 4492–4495.
[40] T.Y. Cheng, M.H. Chen, W.H. Chang, M.Y. Huang, T.W. Wang, Biomaterials 34 (2013) 2005–2016. [41] D. Silva, A. Natalello, B. Sanii, R. Vasita, G. Saracino, R.N. Zuckermann, S.M. Doglia, F. Gelain, Nanoscale. 5 (2013) 704–718. [42] V. Jayawarna, M. Ali, T.A. Jowitt, A.E. Miller, A. Saiani, J.E. Gough, R.V. Ulijn, Adv. Mater. 18 (2006) 611–614. [43] L. Chronopoulou, A.R. Togna, G. Guarguaglini, G. Masci, F. Giammaruco, G.I. Togna, C. Palocci, Soft Matter 8 (2012) 5784–5790. [44] D. Cigognini, D. Silva, S. Paloppi, F. Gelain, J. Biomed. Nanotechnol. 10 (2014) 309–323. [45] P. Moitra, K. Kumar, P. Kondaiah, S. Bhattacharya, Angew. Chem. Int. Edit. 53 (2014) 1113–1117. [46] A. Altunbas, S.J. Lee, S.A. Rajasekaran, J.P. Schneider, D.J. Pochan, Biomaterials 32 (2011) 5906–5914. [47] M. Wu, Z. Ye, Y. Liu, B. Liu, X. Zhao, Mol. BioSyst. 7 (2011) 2040–2047. [48] E. Gatto, A. Quatela, M. Caruso, R. Tagliaferro, M.D. Zotti, F. Formaggio, C. Toniolo, A.D. Carlo, M. Venanzi, ChemPhysChem 15 (2014) 64–68. [49] J.H. Kim, S.Y. Lim, D.H. Nam, J. Ryu, S.H. Ku, C.B. Park, Biosens. Bioelectron. 26 (2011) 1860–1865. [50] R. Orbach, L. Adler-Abramovich, S. Zigerson, I. Mironi-Harpaz, D. Seliktar, E. Gazit, Biomacromolecules 10 (2009) 2646–2651. [51] R.L. Huang, W. Qi, L.B. Feng, R.X. Su, Z.M. He, Soft Matter 7 (2011) 6222–6230. [52] D. Berillo, B. Mattiasson, I.Y. Galaev, H. Kirsebom, J. Colloid Interface Sci. 368 (2012) 226–230. [53] X. Shi, W. Wang, A. Wang, Carbohydr. Polym. 94 (1) (2013) 449–455. [54] J. Wang, C. Liu, Y. Shuai, et al., Colloid Surface B 113 (2014) 223–229. [55] M.B. Hansen, S.E. Nielsen, K. Berg, J. Immunol. Methods 119 (1989) 203–210. [56] A.M. Smith, R.J. Williams, C. Tang, P. Coppo, R.F. Collins, M.L. Turner, A. Saiani, R.V. Ulijn, Adv. Mater. 20 (2008) 37–41. [57] S. Grigoriou, E.K. Johnson, L. Chen, D.J. Adams, T.D. James, P.J. Cameron, 8 (2012) 6788–6791.
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Materials Science and Engineering C journal homepage: www.elsevier.com/locate/msec
Preparation and characterization of a novel sodium alginate incorporated self-assembled Fmoc-FF composite hydrogel Xiao Gong a,b, Christopher Branford-White c, Lei Tao a, Shubai Li d, Jing Quan a, Huali Nie a,b,⁎, Limin Zhu a,⁎⁎ a
College of Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, PR China State Key Laboratory for Modification of Chemical Fibers and Polymer Materials, Donghua University, Shanghai 201620, PR China Institute for Health Research and Policy, London Metropolitan University, London N78 DB, UK d Changzhou Institute of Engineering Technology, Changzhou 213164, PR China b c
a r t i c l e
i n f o
Article history: Received 21 April 2015 Received in revised form 8 August 2015 Accepted 27 August 2015 Available online 2 September 2015 Keywords: Self-assembly Peptide Sodium alginate Hydrogel Structure controllable
a b s t r a c t Dipeptides and their derivatives have attracted tremendous attention owning to their excellent abilities of selfassemble assembling into various structures which have great potentials for applications in biology and/or nanotechnology. In the present study, we dedicate to fabricate a rigid and structure controllable Fmoc-FF/SA composite hydrogel. We found that the modified dipeptide, fluorenyl-9-methoxycarbonyl (Fmoc)-diphenylalanine (Phe-Phe) can self-assemble into rigid hydrogels with structures of nanowires, layered thin films or honeycombs as the change of sodium alginate (SA) concentration. Meanwhile, CD-spectroscopy demonstrated that SA appeared to control the process, but it did not change the arrangement of the Fmoc-FF peptide. Our results demonstrated that the formed hydrogel showed physical and chemical stability as well as possessing good biocompatibility. Rheological measurements showed that the addition of SA could improve the stability of the hydrogel. Cell viability assay revealed that the Fmoc-FF and Fmoc-FF/SA hydrogels are both beneficial for cell proliferation in-vitro. Our results indicated that the fabricated Fmoc-FF/SA composite hydrogels could be used in tissue engineering and drug delivery in the future. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Recently, considerable improvements have been proceeded in the researches of molecular self-assembly systems over the past decades, as is reported, the systems include macromolecules [1,2], proteins [3], nucleic acids [4,5], peptides [6,7], liposomes [8] and graphite oxides [9, 10]. At the same time, peptide-based systems being capable of selfassembling, environment friendly, engaged in molecular recognition and diverse functionality, and have enormous application potential in biological and non-biological fields [11–15]. These molecules usually form hierarchical structures at both nano- and micro-scales. It is assumed that the assembly ordered of these building blocks into defined nanostructures was relied on specific molecular recognition sites. These are facilitated by strong covalent bonds or a combination of week non-covalent interactions, including hydrogen bonds, Vander Waals forces, hydrophobic forces, electro-static interactions, π–π stacking, and chiral dipole–dipole interactions [16,17].
⁎ Correspondence to: H. Nie, College of Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, PR China. ⁎⁎ Corresponding author. E-mail addresses: [email protected] (H. Nie), [email protected] (L. Zhu).
http://dx.doi.org/10.1016/j.msec.2015.08.059 0928-4931/© 2015 Elsevier B.V. All rights reserved.
Peptide self-assembly was first certified by the characterization of the repetitive 16-residue peptide motif n-AEAEAKAKAEAEAKAK-c (EAK 16-II), a fragment of a left-handed Z-DNA binding protein in yeast [18]. After that, more peptide-based materials capable of selfassembling were discovered or synthesized including amphiphilic peptides [19,20], cyclic peptides [21–23], dendritic peptides [6], aromatic dipeptides [24], surfactant-like oligo-peptide [25,26], copolypeptides [27] and the like. Moreover, a class of peptide building blocks sharing common features (RDA16-I, RAD16-II, EAK-I) were designed, and they all showed great potential in facilitating 3D tissue cultures mainly due to the formation of self-assembled nanofiber scaffolds [15]. Among these peptide-based materials, diphenylalanine (FF) and its derivatives have often been used as a model structure for evaluating molecular mechanisms of self-assembly systems. Morphological observations and other approaches have been utilized to investigate the effect of external stimuli on FF self-assembly. These investigations include the effect of role of solvent, assembly surface and pH [28,29]. It was proved that these colloids were not only stimuli-responsive to pH and temperature, but also showed novel and adaptive encapsulation properties [30]. For example, FF-based building blocks are capable of assembling into various functional nano-structures, such as nanotubes [31–34], spherical vesicles [35], nanofibrils and nanoribbons [19,36], nanowires [37,38] and ordered molecular chains [39]. Moreover, peptide hydrogels and their assembly processes have been considered as good candidates
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in biological applications, involving tissue engineering [40–44], drug delivery [45–47], bio-imaging and biosensors [48–50]. Although there have been lots of studies on peptide-based hydrogels, we still knew little about the biological mechanisms of Fmoc-peptide hydrogels, and there were still many shortcomings of the hydrogels in biological safety, cost savings, strength, range of applications and so on. For example, Huang et al. introduced konjac glucomannan (KGM) into the Fmoc-FF system to prepare a potential carrier for drug delivery, but it took nearly 200 min to form a rigid transparent hydrogel [51]. For another example, the frozen glucono-δ-lactone (Gdl) in Fmoc-FF system indeed resulted in the formation of macro-porous hydrogels, however, the cryogels turned mechanically weaker (G′ b 1000 Pa) than the Fmoc-FF hydrogels prepared under the same condition [52]. As a natural polysaccharide, SA is nontoxic, with good solubility and fast gelation propriety, and has been widely applied to the field of food, medicine, etc. Shi et al. prepared a novel and superporous SA-based hydrogel by the self-assembling micelle templating [53]. And KGM/SA hydrogels were prepared using graphene oxide as a drug-binding effector for cancer therapy drug loading and release [54]. In previous studies, scientists introduced polysaccharides into the system to regulate peptide-based hydrogels either from intensity or the structure, but few have shown the capacity to both ensure the strength and the structure of the hydrogels in a short time. The aim of this study is to find a novel and efficient approach to prepare rigid and structure controllable Fmoc-FF/SA composite hydrogels with stable physical and chemical properties. It is hopeful that our study will contribute to understanding the mechanism of peptide selfassembly and the role of polysaccharide in the peptide selfassembly process.
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2. Experimental 2.1. Materials and methods The Fmoc-diphenylalanine (Fmoc-Phe-Phe-OH, Fmoc-FF) peptide in lyophilized form was purchased from Bachem (Bubendorf, Switzerland). Sodium alginate (AR, MW: 398.31) was obtained from Aladdin Chemistry Co. Ltd. Dimethyl sulfoxide (DMSO) was purchased from the Sinopharm Chemical Reagent Co. Ltd. (Shanghai, China). Porcine pepsin (250 units mg−1) and trypsin (3194 units mg−1) were purchased from J&K Scientific Ltd. (Beijing, China). Dulbecco's Modified Eagle Medium (DMEM) was purchased from Genome Biological Technology Co. Ltd. (Hangzhou, China). Penicillin and streptomycin were purchased from Generay Biological Engineering Co. Ltd. (Shanghai, China) and fetal calf serum was purchased from Life Technologies Corporation (America). Other reagents were obtained from the Beijing Chemical Reagent Corporation and were used as received. Water was distilled in a three-stage Millipore Milli-Q plus 185 purification system and had a resistivity N18.2 MΩ. 2.2. Hydrogel preparation SA powders were gradually dispersed in double deionized (dd) H2O (1.0–10.0 mg ml− 1) while stirring, and then stirred for 12 h at room temperature to fully dissolve them. Fmoc-FF stock solution was freshly prepared by dissolving the peptides in DMSO (100.0 mg ml−1) before use. The peptide stock solution was then diluted to a final concentration of 2.0 mg ml−1 by adding dropwise to ddH2O and SA solutions, respectively. The mixture solutions were left at room temperature for three
Fig. 1. A schematic illustration of the gelation process and stability tests of the hydrogel. (a) Molecular structure of Fmoc-FF; (b) white floc emerged after adding peptides; (c) white turbid viscous liquid after sufficient shake; (d) formation of the translucent hydrogel; (e) macroscopic image of the hydrogel samples after ultrasonic processing for 2 h; (f) macroscopic image of the hydrogels after shaking in the shaking bath at 37 °C with 1.0 ml PBS solution; (g) the remained quantity of Fmoc-FF peptide after in vitro digestion of Fmoc-FF hydrogel in simulated gastric fluid; (h) the remained quantity of Fmoc-FF peptide after in vitro digestion of Fmoc-FF hydrogel in simulated intestinal fluid.
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days after sufficient mixing on a miniature oscillator, leading to the formation of the highly transparent Fmoc-FF peptide and Fmoc-FF/SA hybrid hydrogels [6,51]. In order to distinguish the different samples, the Fmoc-FF hydrogel and Fmoc-FF/SA composite hydrogels were labeled 1# (SA, 0 mg ml−1), 2# (SA, 1.0 mg ml−1), 3# (SA, 3.0 mg ml−1), 4# (SA, 5.0 mg ml−1), 5# (SA, 7.0 mg ml−1), and 6# (SA, 10.0 mg ml−1), respectively. 2.3. Hydrogel stability To evaluate the stability of the hydrogels, 0.2 M PBS, pH 7.4 (2.0 ml), containing 0.1% (w/v) NaN3, were added to the upside of isopyknic Fmoc-FF and Fmoc-FF/SA (both 2.0 mg ml−1) hydrogels, respectively. Samples were placed in a rotary shaker at 70 rpm and 37 °C for 12 h and morphological changes were recorded with a digital camera (COOLPIX S80, Nikon Corporation, Japan). 2.4. In vitro digestion The stable hydrogels were immersed in simulated gastric (0.5 ml, 10 mM PBS–HCl, pH 2.0, 20 μg ml−1 pepsin) and simulated intestinal fluids (0.5 ml 10.0 mM PBS, pH 7.4, 20 μg ml−1 trypsin), respectively [51]. The hydrogels were then placed in a rotary shaker, 150 rpm and 37 °C for 24 h. Enzymatic reactions were deactivated by placing samples in boiling water bath for 15 min after centrifugation at 15,000 rpm for 5 min. The supernatants were collected and the amount of Lphenylalanine was determined using the high-performance liquid chromatography (HPLC, Waters, 2489, Waters Corporation, USA) and Agilent TC-C18 column. HPLC was performed at 210 nm using a mobile phase (water: acetonitrile (70:30, v/v) at a flow rate of 1.0 ml min−1, and 20 μl samples were injected. The amount of Fmoc-FF peptide
remained was calculated by removing the amount of L-phenylalanine determined.. 2.5. Rheological measurement In-situ hydrogel formation and rheological properties were measured using an ARES-RFS high rotational rheometer (TA Instruments, USA) at 25 °C with a paralleled plate of geometry (50 mm in diameter, tool inertia 62.5 g·cm2, max auto-tension displacement 3.0 mm, minimum applied dynamic force = 1.0 gmf). Time sweep oscillatory measurements were performed immediately after shaking the samples and preparing mixtures (30 s), diluting the Fmoc-FF stock solutions to give a final concentration of 4.0 mg ml− 1 in ddH2O and SA (1.0 mm gap, fluid density 1.0 g cm−3, strain 0.5%). Whereafter, dyn strain frequency sweep (0.1–100 Hz, strain 0.5%) measurement was conducted to determine the frequency dependent storage and loss sheer moduli, where in the time sweep oscillatory test was performed [51,52]. All tests were repeated three times and the average value was taken. 2.6. Scanning electron microscopy (SEM) To gain insight into the real morphology of the self-assembled structures of the (composite) hydrogels, all the samples (3 days old) were prepared by rapid freezing for 2 h at − 80 °C, and drying for 24 h in a freeze drier (Christ Alpha1-2, Osterode, Germany). In order to observe the self-assembled morphology of Fmoc-FF in the composite hydrogels, the samples were dispersed in water and SA was removed by membrane filtration. All the samples were sputter-coated and then images were recorded using a JSM-5600LV scanning electron microscope (JEOL, Tokyo, Japan).
Fig. 2. Dynamic rheology analysis of self-assembled Fmoc-FF hydrogel (1#) and Fmoc-FF/SA composite hydrogel (2#). (a) Time dependence of G′ and G″ of Fmoc-FF peptide hydrogel; (b) time dependence of G′ and G″ of Fmoc-FF/SA composite hydrogel; (c) the change in G′ and G″ of Fmoc-FF peptide hydrogel with the increased frequency (0.1–100 Hz); (d) the change in G′ and G″ of Fmoc-FF/SA composite hydrogel with the increased frequency (0.1–100 Hz).
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2.7. Transmission electron microscopy (TEM)
2.10. Circular dichroism spectroscopy (CD)
After freezing–drying, Fmoc-FF and Fmoc-FF/SA xerogels were negatively stained with phosphotungstic acid (1%) for 2 min. Then a drop of the sample xerogel was placed on a 400-mesh copper grid and dried at room temperature. Samples were viewed at 80 kV using a JEM-2100 electron microscope (JEOL Ltd., Japan).
CD spectra were recorded using a Jasco J-810 CD spectrometer (Jasco, Glasgow, UK), 190–260 nm, with a resolution of 0.5 nm at room temperature. Samples were prepared by dispersing the xerogels in aqueous solutions to avoid DMSO interfacing. The path length of the quartz cell was 1.0 mm and 0.2 mg ml− 1 [51,52]. All tests were repeated three times and the average value was taken.
2.8. Thermogravimetric analysis (TGA) 2.11. Viability analysis TGA (TA 2050, America) was conducted after the freezing–drying process. The samples were heated at the rate of 8 °C min− 1 under nitrogen. 2.9. Fourier transform infrared spectroscopy (FTIR) FTIR spectra of Fmoc-FF/SA xerogels, Fmoc-FF and SA were undertaken on a Nicolet-6700 FTIR spectrometer (Thermo Fisher, USA) over 3750–600 cm−1 at a resolution of 2 cm−1; 32 scans were taken.
Studies on the scaffold effects of the composite hydrogels for tissueengineering applications and in-vitro cellular experiments were undertaken using the following protocol. Fresh Fmoc-FF and Fmoc-FF/SA composite hydrogels (300 μl) were prepared in water and SA, respectively. After autoclaving, samples were placed in a 24- hole culture plate system (Corning) and then fumigated with alcohol overnight. For cell-growth pig iliac endothelial cells (PIEC) were grown in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10%
Fig. 3. SEM images of self-assembled Fmoc-FF peptide hydrogel and Fmoc-FF/SA composite hydrogels. (a) Fmoc-FF peptide hydrogel; (b)–(f) Fmoc-FF/SA composite hydrogels with different concentrations of sodium alginate (1.0, 3.0, 5.0, 7.0, 10.0 mg ml−1, respectively).
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fetal calf serum, 100 U ml−1 penicillin and 100 U ml−1 streptomycin. Cells were maintained at 37 °C in a humidified atmosphere containing 5% CO2. Subconfluent cells were harvested by trypsinization, counted, and diluted in the cells media to 1.0 × 106 cells ml−1. Before cell implantation, the hydrogels were washed (three times) with PBS (pH 7.4, 0.01 M) and DMEM, respectively. Then 20 μl of PIEC (2 × 104 cells) was over laid on the hydrogels. After 24 h incubation at 37 °C with 5% CO2 in the constant temperature incubator, the viability of the cells was determined using an MTT assay [6,55]. The cell medium was replaced daily and the results from triplicate samples taken. To qualitatively confirm cell viability and proliferation upside the hydrogels, confocal laser scanning microscopy (CLSM, Carl Zeiss LSM 700, Germany) was used. Cover slips with a diameter of 14 mm were pretreated with 5% HCl, 30% HNO3, and 75% alcohol and then fixed in 24-well tissue culture plate. Total number of 2 × 104 PIEC was seeded per well upside the hydrogels and cultivated for 7 days. Then the cells were rinsed with PBS for 3 times fixed with glutaraldehyde (2.5%) for
15 min at 4 °C, and stained with rhodamine (1 μg ml−1) for 15 min at 37 °C using a standard procedure. Finally, all samples were washed with PBS and observed via CLSM. 3. Results and discussion 3.1. Hydrogel preparation The Fmoc-FF hydrogel was prepared according to the following method: First, dissolve the Fmoc-FF peptide in DMSO (Fig. 1a), then inject it into ddH2O (Fig. 1b). Meanwhile, the Fmoc-FF turned into white floc immediately (Fig. 1c). Whereafter, shake the sample tube to mix evenly. Within a few minutes, a stable transparent hydrogel formed (Fig. 1d) from a state of white turbid colloid. The Fmoc-FF/SA composite hydrogel was prepared by mixing the peptide with SA at different concentrations. The gelation of the Fmoc-FF was also accompanied by the formation of a transparent hydrogel, though it took a little longer time
Fig. 4. TEM and SEM micrographs of self-assembled Fmoc-FF peptide hydrogel and Fmoc-FF/SA hydrogels. (a) TEM micrograph of Fmoc-FF; (b)–(d) TEM micrographs of Fmoc-FF/SA composite hydrogels (3#, 5#, 6#); (e) SEM image of stacked massive structure of Fmoc-FF peptide; (f) SEM image of membranous Fmoc-FF structure after removal of SA (5#).
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3.3. Rheology characterization
Fig. 5. TG curves of (a) the dried sodium alginate; (b) Fmoc-FF/SA composite hydrogel (2#); and (c) Fmoc-FF.
(ranging from several minutes to several quarters) to form a stable and transparent composite hydrogel as the concentration of sodium alginate increased.
3.2. Stability tests The stability of the hydrogels was assessed by ultrasonic concussing and incubation at pH 7.4 and 37 °C. The change of the two hydrogel samples was noted after they had been treated by ultrasonication for 2 h. It was clear that both the Fmoc-FF hydrogel (1#) and Fmoc-FF/SA composite hydrogels (2#–6#) remained almost unchanged (Fig. 1e). As the concentration of SA increased, the state of hydrogel systems gradually changed from turbid viscous fluid to samples with clear “water layer–gel layer” interface (Fig. 1f), showing preferable properties of anti-phosphate buffer. These results also illustrated that the interference of SA enhanced the stability of the hydrogel system. In the presence of proteases (pepsin and trypsin), in-vitro digestion of the FmocFF peptide and the composite hydrogels were evaluated. No detection of phenylalanine was found after 24 h and it happened because these structures were resistant to proteolytic attack (Fig. 1g and Fig. 1h). The results indicated that this system could probably provide as a platform for future applications of oral drugs and their delivery [6].
The mechanical properties of the hydrogels are important for assessing their suitability for future applications. Thus dynamic rheology experiment was undertaken to investigate the kinetic formation of hydrogel and the effect of SA on the gelation process. Fig. 2a showed the storage modulus (G′) and shear modulus (G″) change over time. G ′ and G″ were almost invariable within 60 s. However, from 60 s to 150 s, the values both rose sharply and reached a plateau at about 150 s. From the date, the storage modulus G′ (~ 4300 Pa) exceeded that of the loss modulus G″ (~400 Pa) by ~11 times, indicating the formation of a steady viscoelastic hydrogel [37,51]. However, the gelation time (i.e. the time after which G′ becomes larger than G″) of Fmoc-FF/ SA (2#) was much shorter than the Fmoc-FF hydrogel and was even difficult to record (Fig. 2b), though the stable Fmoc-FF/SA composite hydrogel formed only with a G′ of 2000 Pa, and G″ of 200 Pa. This result was also consistent with the findings that the rigid transparent FmocFF hydrogel formed within a few minutes, but a longer time for the Fmoc-FF/SA composite hydrogel. It is assumed that the steric hindrance of SA, and strong hydrogen bonds between the molecules, impeded the movement of Fmoc-FF molecules and their aggregation, thereby, prolonged the process of self-assembly. Nondestructive frequency sweep indicated that the shear modulus G ″ of Fmoc-FF dramatically decreased with frequency, especially when the frequency increased to 30 Hz, despite a slight increase of G′ for Fmoc-FF hydrogel between 0.1 Hz to 100 Hz (Fig. 2c). However, in contrast with Fmoc-FF hydrogel, both G′ and G″ of the Fmoc-FF/SA composite hydrogel were not affected by the increase of shear frequency (Fig. 2d). In short, the composite hydrogel (SA, 1.0 mg ml−1) exhibited better frequency stability and toughness than the Fmoc-FF hydrogel within the frequency range. That is because SA contains a large number of hydroxyl groups, making it easy to form intermolecular hydrogen bond. Accordingly, the enhanced hydrogen bonding among selfassembled peptide, water and SA (–OH groups) strengthen the stability of composite hydrogels.
3.4. Morphology characterization The morphological structures of the self-assembled hydrogels were observed by SEM and TEM, respectively. As shown in Fig. 3a, when the Fmoc-FF stock solution was directly injected into ddH2O, selfassembled blocky structure in micro-scale was formed, which differed from previous reports [6,24]. However, when SA (1.0 mg ml−1) was introduced into the Fmoc-FF system, the morphology of the assembled
Fig. 6. (a) FTIR spectra of Fmoc-FF hydrogel, sodium alginate and Fmoc-FF/SA composite hydrogel (2#); (b) CD spectra of Fmoc-FF peptide and Fmoc-FF/SA composite hydrogel (2#).
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structure, it was comprehensible that the Fmoc-FF hydrogel was the toughest among the all of hydrogels. The Fmoc-FF hydrogel finally self-assembled into a micro-sized blocky structure, while the Fmoc-FF/ SA composite hydrogels formed nanowire networks or “honeycomblike” structure etc. The presence of numerous cavities, perhaps contributed to the decrease of G′. TEM showed that Fmoc-FF hydrogel consisted of plenty of blocky structures (Fig. 4a, c) in similar morphology. Fmoc-FF in the composite hydrogel (3#) can self-assemble into long nanowires which are several microns in length and several tens of nanometers in width (Fig. 4b). The “honeycomb sheet” structure of the composite hydrogel (6#) was also confirmed by TEM (Fig. 4d). The reason why we did not observe a complete three-dimensional cellular structure was that the material was redissolved prior to TEM observation. Compared with the membranous structure of the composite hydrogel (5#, Fig. 4c), the “honeycomb sheet” was thinner and flexible with clear wrinkles on the surface. All these morphological observations illustrated that the SA played more important roles as an inducer or a template to immobilize Fmoc-FF.. Fig. 7. Cell viability tests on the surface of Fmoc-FF hydrogel and Fmoc-FF/SA composite hydrogels for 1, 3, 5, and 7 days.
hydrogel changed from blocky structure to short nanowires, tens of nanometers to several microns in length (Fig. 3b). Moreover, when we increased the concentration of SA to 3.0 mg ml−1, nanowires from several to tens of microns in length appeared (Fig. 3c). To further increase the concentration of SA (5.0 mg ml− 1), the nanowire structure was hardly observed and films containing a large number of folds generated (Fig. 3d). As shown in Fig. 3e, when SA of 7.0 mg ml−1 was applied, pieces of thin films constituted the entire hydrogel scaffold. Fig. 3f revealed that in the presence of high concentration of SA (10.0 mg ml−1), the composite hydrogels self-assembled into “honeycomb-like” structure that contained several cavities. It was reasonable to assume that the removal of water held within the honeycomb scaffold contributed to the structural stability and so shrank the “honeycomb sheet” to form a “shriveled and complicated” pile. Various structures were formed by controlling the concentration of SA, and this could be due to the steric hindrance of SA and the impact of strong non-covalent bond formation that facilitated Fmoc-FF assembly after aggregation, such as the strong hydrophobic interaction of the polypeptide, π–π stacking of the Fmoc-FF molecules and hydrogen-bond interaction with SA molecule [25,28]. From the perspective of hydrogel
3.5. Supramolecular arrangement TGA was performed to investigate the combination of Fmoc-FF and SA. As shown in Fig. 5a, the thermal decomposition of sodium alginate was divided into three stages (named Stages 1, 2, 3). Stage 1 (within 100 °C) was the effect of lactonization and transglycosylation; Stage 2 (from 220 to 250 °C) was fracture of the sodium alginate skeleton, and the final stage was the carbonization effect. For Fmoc-FF, only one stage of weight loss (from 220 to 300 °C) occurred, and this accounted for 90% weight loss (Fig. 5c). It was clear in Fig. 5b that the weight loss of Fmoc-FF/SA from 200 to 400 °C compromised that for sample SA and Fmoc-FF, indicating that the composite hydrogel sample integrated the properties of the two materials. FTIR was used to verify the above assumption. The spectrum of the Fmoc-FF/SA xerogel after freezing–drying (Fig. 6a) showed the position of the amide I band, a weak peak at 1692 cm−1 and a stronger one at 1652 cm−1, indicating the predominant β-sheet character [19]. It was clear that Fmoc-FF/SA composite hydrogel showed typical amide II and amide III adsorption peak centered at 1538 cm−1 and 1258 cm−1 that attributed to Fmoc-FF and carboxylate adsorption peak at 1606 cm−1 and 1414 cm−1 that attributed to SA. The presence of all these characteristic peaks confirmed a good blend of SA and Fmoc-FF,
Fig. 8. Confocal microscopic images of (a1–a3) Rhodamine stained PIEC upside of Fmoc-FF hydrogel; (b1–b3) Rhodamine stained PIEC upside of Fmoc-FF/SA composite hydrogel (2#).
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which laid a foundation for regulating the structure of self-assembled peptide without changing the arrangement of the Fmoc-FF. The secondary molecular structure of Fmoc-FF peptide was assessed using circular dichroism (CD). The spectra of Fmoc-FF peptide (Fig. 6b) were similar to that of the composite hydrogel, with positive peaks at ~ 194 nm, and ~ 214 nm, indicating the β-sheet arrangement of the Fmoc-FF molecular structure [24,34]. Random coil conformation of the Fmoc-FF molecular structure could be confirmed by a strong negative peak at 197 nm and a weak positive wide peaks at 217–223 nm. For the Fmoc-FF/SA composite hydrogel, the strong peak at ~198 nm, and the negative peak at ~215 nm demonstrated the β-sheets conformation, while the negative peak at 201 nm and the positive peak ~219 nm were due to random coiling [56]. Meanwhile, morphology of Fmoc-FF in Fig. 6b was also a good explanation of the role SA played: SA served as an inducer to assemble the composite hydrogel and a template to immobilize the Fmoc-FF. Previous studies reported that molecules with hydroxyl, benzene and other functional groups were capable of interacting with peptide molecules to form different self-assembled materials with various shapes and functions [48,57]. It is well known that SA owns a large numbers of hydroxyl groups, which would certainly influence the arrangement of the Fmoc-FF peptide due to the enhanced hydrogen bond among SA, water, and peptide. To study the effect of SA on the self-assembly of Fmoc-FF molecular, SA was removed from the xerogels by dissolving in ddH2O and vacuum filtration. It was confirmed that Fmoc-FF and SA have been mixed well together. If there is some interaction between Fmoc-FF and SA, the structure of the Fmoc-FF/SA composite hydrogel should be seriously damaged with the dissolution of SA. If SA just played a role of regulating agent, the self-assembled structures might be well preserved. However, compared with those without being washed, the morphology of selfassembled Fmoc-FF remained the same after the removal of SA (Fig. 4e, f). This phenomenon directly indicated that SA played an important role in “inducing agent” during the Fmoc-FF self-assembly process and didn't change the arrangement of the Fmoc-FF peptide [51]. Based on the above research, we can summarize that the role of SA in the Fmoc-FF/SA system mainly involves: (i) serving as an inducer to form the composite hydrogel and a template to immobilize Fmoc-FF; (ii) contributing to Fmoc-FF aggregation in situ and growing into desired structures through the steric hindrance of SA under the different concentration; (iii) stabilizing the 3D gel network structure due to the enhanced hydrogen bonding among self-assembled peptide, water and SA (–OH groups). 3.6. Cell viability assay To assess and evaluate the ability of composite hydrogels to be used in biological applications, cell viability was analyzed following 1, 3, 5, and 7 days of incubation period (Fig. 7). After one day, cells were taken from all peptide hydrogel surfaces, and they all increased noticeably, reaching nearly 50% growth rate compared with the control. After 3 days, the number of cells on the composite hydrogels with SA (1.0 mg ml−1–10.0 mg ml−1) was nearly identical to that Fmoc-FF hydrogel without SA (1#), which increased by ~ 77% compared with the control experiment, showing good cell viability. Over the next few days, cells continued to grow. Particularly, cells on the Fmoc-FF peptide hydrogel proliferated by 93% after 7 days of cultivation compared with the same stage of control. However there was a small decrease in growth rate of cells on the composite hydrogels after days 5–7, with the cell population increase of 70% compared with the control group. Therefore, viability assay showed that sodium alginate had little effect on cell growth. The survival status of PIEC cultured on the hydrogels was qualitatively confirmed via confocal microscopic imaging of the red fluorescence of rhodamine. As shown in Fig. 8a1–a3, when the Fmoc-FF hydrogel was inoculated with PIEC, cells grew rapidly and were fairly homogenous on the surface. Fluorescent photomicrographs also
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showed good morphology of PIEC in the composite hydrogel (Fig. 8b1–b3). By comparing Fig. 8b2 with Fig. 8b1 and b3, it was clear that cells grew not only on the surface of the composite hydrogel, but also in the internal space of the nanowire networks as the red arrows pointed to. It was also consistent with Fig. 7. In short, Fmoc-FF finally self-assembled into blocky structures in micro-scale, while the FmocFF/SA composite hydrogels formed nanowire networks or “honeycomb-like” structure, leading to the weaker of G′. However, from the perspective of applications, especially in tissue engineering as well as cell and/or tissue regeneration, Fmoc-FF/SA hydrogels with rich cavity structure could provide the cells adequate growth space. 4. Conclusions In this study, we designed a novel composite hydrogel by inducing SA into Fmoc-FF peptide system. The hydrogel was stable in buffer and proteolytic digestion. Rheological results revealed that the asprepared hydrogels were consistently intact and the morphological observations indicated that the Fmoc-FF/SA can self-assemble into nanowires, layered thin films, or honeycomb-like structures, while the Fmoc-FF peptide self-assembled into blocky structure. The driving force for the self-assembled nanostructure was inferred to be due to hydrogen bonding and aromatic packing factors. SA also played an important role in regulating the movement and combination of Fmoc-FF molecules during assembly. Data from CD implied that the hydrogels had similarities in terms of β-sheet and random coil peptides. Finally, cell viability assay demonstrated that Fmoc-FF/SA composite hydrogels exhibited good biocompatibilities and enhanced the growth of epithelial cells. It is expected that the novel composite hydrogels would be utilized in various broad applications in future tissue engineering as well as cell and/or tissue regeneration. Acknowledgments The project was funded by the Shanghai Natural Science Foundation (14ZR1401300), National Key Fund (EG2014021), Qing Lan Project, Changzhou Applied Science Foundation (CJ20120014) and State Key Laboratory for Modification of Chemical Fibers and Polymer Materials Donghua University. References [1] B.M. Blunden, H.X. Lu, M.H. Stenzel, Biomacromolecules 14 (2013) 4177–4188. [2] W.I. Hung, C.H. Chang, Y.H. Chang, P.S. Wu, C.B. Hung, K.C. Chang, M.C. Lai, S.C. Hsu, Y. Wei, X.R. Jia, J.M. Yeh, Langmuir 29 (2013) 12075–12083. [3] Y.P. Chuan, Y.Y. Fan, L.H.L. Lua, A.P.J. Middelberg, J. R. Soc. Interface 7 (2010) 409–421. [4] P.W.K. Rothemund, A. Ekani-Nkodo, N. Papadakis, A. Kumar, J. Am.Chem.Soc. 126 (2004) 16344–16352. [5] A. Gangar, A. Fegan, S.C. Kumarapperuma, P. Huynh, A. Benyumov, C.R. Wagner, Mol. Pharm. 10 (2013) 3514–3518. [6] A. Mahler, M. Reches, M. Rechter, S. Cohen, E. Rigid Gazit, Adv, Mater 18 (2006) 1365–1370. [7] N. Nakatsuka, S.N. Barnaby, K.R. Fath, I.A. Banerjee, J Biomat Sci-Polym E. 23 (2012) 1843–1862. [8] D.G. Yu, C. Branford-White, G.R. Williams, S.W.A. Bligh, K. White, L.M. Zhu, N.P. Chatterton, Soft Matter 18 (2006) 1365–1370. [9] C.M. Chen, Q.H. Yang, Y.G. Yang, W. Lv, Y.F. Wen, P.X. Hou, M.Z. Wang, H.M. Cheng, Adv. Mater. 21 (2009) 3007–3011. [10] C. Wu, X.Y. Huang, G.L. Wang, L.B. Lv, G. Chen, G.Y. Li, P.K. Jiang, Adv. Funct. Mater. 23 (2013) 506–513. [11] L. Liu, X. Liu, H. Deng, Z. Wu, J. Zhang, B. Cen, Q. Xu, A. Ji, J Biomat Sci-Polym E. 25 (2014) 1331–1345. [12] D. Silva, A. Natalello, B. Sanii, R. Vasita, G. Saracino, R.N. Zuckermann, S.M. Doglia, F. Gelain, Nanoscale 23 (2013) 506–513. [13] P. Kongsuphol, S.K. Arya, C.C. Wong, L.J. Polla, M.K. Park, Biosens. Bioelectron. 55 (2014) 26–31. [14] P. Moitra, K. Kumar, P. Kondaiah, S. Bhattacharya, Angew. Chem. Int. Ed. 53 (2014) 1113. [15] S.G. Zhang, Nat. Biotechnol. 53 (2014) 1113–1117. [16] J.E. Huheey, E.A. Keiter, R.L. Keiter, Inorganic Chemistry: Principles of Structure and Reactivity, Prentice Hall, Upper Saddle River, NJ, 1997. [17] P.Y. Bruice, Organic Chemistry, Prentice Hall, Upper Saddle River, NJ, 2003.
486
X. Gong et al. / Materials Science and Engineering C 58 (2016) 478–486
[18] S.G. Zhang, T. Holmes, C. Lockshin, A. Rich, P. Natl. Acad. Sci. USA 90 (1993) 3334–3338. [19] Y.Y. Lin, Y. Qiao, P.F. Tang, Z.B. Li, J.B. Huang, Soft Matter 7 (2011) 2762–2769. [20] J.D. Hartgerink, E. Beniash, S.I. Stupp, Science 294 (2001) 1684–1688. [21] M.R. Ghadiri, J.R. Granja, R.A. Milligan, D.E. Mcree, N. Khazanovich, Nature 366 (1993) 324–327. [22] L. Ruiz, S. Keten, J. Eng. Mech. 140 (2014) 431–442. [23] Y.Z. Wang, S.J. Yi, L.M. Sun, Y.J. Huang, S.C. Lenaghan, M.J. Zhang, J. Biomed. Nanotechnol. 10 (2014) 445–454. [24] M. Hughes, P.W.J.M. Frederix, J. Raeburn, L.S. Birchall, J. Sadownik, F.C. Coomer, I.H. Lin, E.J. Cussen, N.T. Hunt, T. Tuttle, S.J. Webb, D.J. Adams, R.V. Ulijn, Soft Matter 8 (2012) 5595–5602. [25] X.J. Zhao, S.G. Zhang, Chem. Soc. Rev. 35 (2006) 1105–1110. [26] Y.Y. Lin, T. Qiao, P.F. Tang, Z.B. Li, J.B. Huang, Soft Matter 7 (2011) 2762–2769. [27] T.J. Sanborn, P.B. Messersmith, A.E. Barron, Biomaterial. 23 (2002) 2703–2710. [28] X.H. Yan, Y. Cui, Q. He, K.W. Wang, J.B. Li, Chem. Mater. 20 (2008) 1522–1526. [29] R.L. Huang, W. Qi, R. Su, Soft Matter 7 (2011) 6418–6421. [30] X.H. Yan, P.L. Zhu, J.B. Fei, J.B. Li, Adv. Mater. 22 (2010) 1283–1287. [31] M. Reches, E. Gazit, Nat. Nanotechnol. 1 (2006) 195–200. [32] R.J.A. Hill, V.L. Sedman, S. Allen, P.M. Williams, M. Paoli, L. Adler-Abramovich, E. Gazit, L. Eaves, S.J.B. Tendler, Adv. Mater. 19 (2007) 4474–4479. [33] X.H. Yan, Y. Cui, Q. He, K.W. Wang, J.B. Li, W.H. Mu, B.L. Wang, Z.C. Qu-yang, ChemEur J 14 (2008) 5974–5980. [34] P. Bharagava, Y.F. Tu, J.X. Zheng, H.M. Xiong, R.P. Quirk, S.Z.D. Cheng, J. Am. Chem. Soc. 129 (2007) 1113–1121. [35] M. Reches, E. Gazit, Nano Lett. 4 (2004) 581–585. [36] I. Cherny, E. Gazit, Angew. Chem. Int. Edit. 47 (2008) 4062–4069. [37] T.H. Han, J. Kim, J.S. Park, C.B. Park, H. Ihee, S.O. Kim, Adv. Mater. 19 (2007) 3924–3927. [38] J. Ryu, C.B. Park, Adv. Mater. 20 (2008) 3754–3758. [39] M. Lingenfelder, G. Tomba, G. Costantini, L.C. Ciacchi, A. De Vita, K. Kern, Angew. Chem. Int. Ed. 46 (2007) 4492–4495.
[40] T.Y. Cheng, M.H. Chen, W.H. Chang, M.Y. Huang, T.W. Wang, Biomaterials 34 (2013) 2005–2016. [41] D. Silva, A. Natalello, B. Sanii, R. Vasita, G. Saracino, R.N. Zuckermann, S.M. Doglia, F. Gelain, Nanoscale. 5 (2013) 704–718. [42] V. Jayawarna, M. Ali, T.A. Jowitt, A.E. Miller, A. Saiani, J.E. Gough, R.V. Ulijn, Adv. Mater. 18 (2006) 611–614. [43] L. Chronopoulou, A.R. Togna, G. Guarguaglini, G. Masci, F. Giammaruco, G.I. Togna, C. Palocci, Soft Matter 8 (2012) 5784–5790. [44] D. Cigognini, D. Silva, S. Paloppi, F. Gelain, J. Biomed. Nanotechnol. 10 (2014) 309–323. [45] P. Moitra, K. Kumar, P. Kondaiah, S. Bhattacharya, Angew. Chem. Int. Edit. 53 (2014) 1113–1117. [46] A. Altunbas, S.J. Lee, S.A. Rajasekaran, J.P. Schneider, D.J. Pochan, Biomaterials 32 (2011) 5906–5914. [47] M. Wu, Z. Ye, Y. Liu, B. Liu, X. Zhao, Mol. BioSyst. 7 (2011) 2040–2047. [48] E. Gatto, A. Quatela, M. Caruso, R. Tagliaferro, M.D. Zotti, F. Formaggio, C. Toniolo, A.D. Carlo, M. Venanzi, ChemPhysChem 15 (2014) 64–68. [49] J.H. Kim, S.Y. Lim, D.H. Nam, J. Ryu, S.H. Ku, C.B. Park, Biosens. Bioelectron. 26 (2011) 1860–1865. [50] R. Orbach, L. Adler-Abramovich, S. Zigerson, I. Mironi-Harpaz, D. Seliktar, E. Gazit, Biomacromolecules 10 (2009) 2646–2651. [51] R.L. Huang, W. Qi, L.B. Feng, R.X. Su, Z.M. He, Soft Matter 7 (2011) 6222–6230. [52] D. Berillo, B. Mattiasson, I.Y. Galaev, H. Kirsebom, J. Colloid Interface Sci. 368 (2012) 226–230. [53] X. Shi, W. Wang, A. Wang, Carbohydr. Polym. 94 (1) (2013) 449–455. [54] J. Wang, C. Liu, Y. Shuai, et al., Colloid Surface B 113 (2014) 223–229. [55] M.B. Hansen, S.E. Nielsen, K. Berg, J. Immunol. Methods 119 (1989) 203–210. [56] A.M. Smith, R.J. Williams, C. Tang, P. Coppo, R.F. Collins, M.L. Turner, A. Saiani, R.V. Ulijn, Adv. Mater. 20 (2008) 37–41. [57] S. Grigoriou, E.K. Johnson, L. Chen, D.J. Adams, T.D. James, P.J. Cameron, 8 (2012) 6788–6791.