High-Strength Films Consisted of Oriented Chitosan Nanofibers for

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High-Strength Films Consisted of Oriented Chitosan Nanofibers for Guiding Cell Growth Kunkun Zhu, Jiangjiang Duan, Jinhua Guo, Shuangquan Wu, Ang Lu, and Lina Zhang Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.7b00936 • Publication Date (Web): 09 Oct 2017 Downloaded from http://pubs.acs.org on October 11, 2017

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High-Strength Films Consisted of Oriented Chitosan Nanofibers for Guiding Cell Growth

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Kunkun Zhu, Jiangjiang Duan, Jinhua Guo, Shuangquan Wu, Ang Lu*, Lina Zhang*

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College of Chemistry and Molecular Sciences, Wuhan University, Wuhan 430072, China

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*Correspondence to: [email protected] (L. Zhang), [email protected] (A. Lu)

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ACS Paragon Plus Environment

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ABSTRACT

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Chitosan has biocompatibility and biodegradability, however the practical use of the bulk

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chitosan materials is hampered by its poor strength, which can not satisfy the mechanical

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property requirement of organs. Thus, the construction of highly strong chitosan-based

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materials has attracted much attention. Herein, the high strength nanofibrous hydrogels and

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films (CS-E) were fabricated from the chitosan solution in LiOH/KOH/urea aqueous system

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via a mild regenerating process. Under the mild condition (ethanol at low temperature)

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without the severe fluctuation in the system, the alkaline-urea shell around the chitosan chains

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was destroyed, and the naked chitosan molecules had sufficient time for the orderly

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arrangement in parallel manner to form relatively perfect nanofibers. The nanofibers

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physically cross-linked to form CS-E hydrogels, which could be easily oriented by drawing to

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achieve a maximum orientation index of 84 %, supported by the scanning electron

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microscopy and two-dimensional wide-angle X-ray diffraction. The dried CS-E films could be

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bent and folded arbitrarily to various complex patterns and shapes. The oriented CS-E films

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displayed even ultra-high tensile strength (282 MPa), which was 5.6 times higher than the

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chitosan films prepared by the traditional acid dissolving method. The CS-E hydrogels

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possessed hierarchically porous structure, beneficial to the cell adhesion, transportation of

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nutrients, and removal of metabolic byproducts. The cell assay results demonstrated that the

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CS-E hydrogels were no cytotoxicity, and osteoblastic cells could adhere, spread, and

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proliferate well on their surface. Furthermore, the oriented CS-E hydrogels could regulate the

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directional growth of osteoblastic cells along the orientation direction, on the basis of the

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filopodia of the cells to extend and adhere on the nanofibers. This work provided a novel 2


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approach to construct the oriented high strength chitosan hydrogels and films.

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Keywords: chitosan film, nanofiber, mild regenerating process, high orientation, cell

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proliferate

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INTRODUCTION

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Biological macromolecules with excellent biocompatibility, bioactivity and non-toxicity

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are good candidates for biomedical materials.1-3 Chitin is the second most abundant naturally

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occurring polymer and exists mainly in the exoskeletons of crabs and shrimps.4 When the

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degree of deacetylation of chitin is more than about 60%, it becomes alkaline polysaccharide

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with the –NH2 groups, which is called as chitosan. Chitosan is readily soluble in dilute acidic

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solutions, and bulk materials can be constructed in non-solvents via regeneration.5-7 Due to

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the intrinsic biocompatibility, biodegradability, strong affinity, nontoxicity, antimicrobial

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activity, and low immunogenicity, chitosan based materials are considered to be potential in

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the biomaterial fields.8-11 However, the practical use of the bulk chitosan materials is

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hampered by its poor strength, which can not satisfy the mechanical property requirement of

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organs, such as cortical bone (longitudinal, ~130-190 MPa; cross, ~40-60 MPa), trabecular

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bone (2-7 MPa), dentine (21-53 MPa), human Achilles tendon (71-86 MPa), and wet compact

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bone (124-174 MPa).12-16 Thus, the good mechanical properties is the basic requirements for

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such applications. In the last decades, strategies including nanocomposite, blending with

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polymers, chemical crosslinking have been developed to improve the mechanical properties

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of chitosan films.17 Unfortunately, such techniques usually result in only a moderate 3


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enhancement of mechanical strength but sometimes sacrifice the biocompatibility and

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biodegradability.18 Therefore, it still remains as a great challenge to enhance the mechanical

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properties of the bulk chitosan materials, retaining the unique bioactive features of chitosan

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macromolecules. Furthermore, the tensile orientation requires material with strong strength to

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achieve more functionality.

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Recently, the attractive advancement of nanoscience and nanotechnology are witnessed,

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and the nanoscale structures significantly promotes the evolution of the functional materials in

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various fields. For example, nanofibrous materials with good mechanical strength, high light

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transmittance, large surface area, three-dimensional structure and orientation properties are

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widely applied in optical devices, filtration, battery, food preservation, effluent treatment,

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biomedical and tissue engineering.19-23 To generate such nanostructure, the “bottom-up”

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approach is frequently utilized, via which molecules assemble to form nanofibers and

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nanofibrous structure.24-26 By introducing nanofibrous structure, the mechanical strength of

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the bulk material from cellulose, chitin, etc. were greatly improved.27-29 In our laboratory,

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chitosan hydrogels with excellent mechanical properties have been fabricated in the

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alkaline/urea aqueous system, and the compression fracture stress of the chitosan hydrogels is

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nearly 100 times that of the chitosan hydrogels prepared by the traditional acid dissolving

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method, confirming that the nanofibrous network microstructures contribute greatly to the

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reinforcement.30, 31 However, the chitosan films with excellent mechanical properties have

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been scarcely reported.

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Herein, a novel approach to directly prepare chitosan films with high strength was

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developed, for the first time. Our strategy was to slow the self-aggregation rate of the 4


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macromolecules for their orderly rearrangement, namely in a mild regenerated condition (low

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temperature) to form perfect nanofibrous structure. Briefly, the chitosan solution in

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LiOH/KOH/urea aqueous system was regenerated in ethanol at low temperature, in which the

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alkaline-urea shell around the chitosan chains was destroyed. Thus, under this mild

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environment without the severe fluctuation (including heat-induced Brownian motion, violent

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collision between molecules, rapid diffusion and exchange between solvent/non-solvent) in

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the system, the naked chitosan molecules could align sufficiently in parallel manner to form

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relatively perfect nanofibers, which subsequently were physically cross-linked to form

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networks. The chitosan hydrogels were further oriented by drawing, and the resultant dry

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films exhibited good mechanical properties and orientation structure. Furthermore, the

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oriented hydrogels are capable of regulating the directional growth of cells.

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EXPERIMENTAL SECTION

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Materials. Chitosan (CS, commercial grade) with weight-average molecular weight of

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36.0×104 was purchased from Ruji Biotechnology Co., Ltd. (Shanghai, China). The degree of

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deacetylation (DD) of CS was 89% determined by two-abrupt-change potentiometric titration

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method and calculated using the follow equation30

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α =

× × × × .

× 100%

(1)

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Where CNaOH and ∆v are the concentration and volume of NaOH consumption between the

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two abrupt changes of pH, respectively, m is the dry weight of a chitosan sample, and α is the

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degree of deacetylation of the chitosan sample. Other chemical reagents were purchased from

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Sinopharm chemical Reagent Co., China, and used without further purifications. 5


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Preparation of chitosan films. The aqueous solution containing LiOH/KOH/urea/H2O of

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4.5:7:8:80.5 by weight was used as solvent of chitosan. The chitosan powders were dispersed

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in the solvent with stirring for 1 min, and then stored at -30 oC for 6 h until completely frozen.

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Then, the frozen solid was fully thawed and stirred extensively at room temperature. After

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removing air bubbles by centrifugation at 5 oC, a transparent chitosan solution with

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concentration of 4 wt% or 6 wt% was obtained. Subsequently, the chitosan solution was

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spread on a glass plate and then immersed in ethanol at low temperature to get the chitosan

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hydrogels with thickness of 0.9 mm. The resulted hydrogels were washed with distilled water

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to remove any residuals, and then fixed on a glass plate to air-dry at ambient temperature to

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form dry films (with thickness of 0.15 mm), coded as CS-E-20, CS-E-15, CS-E-10, CS-E-5,

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CS-E0, CS-E30, corresponding to regeneration temperatures of -20 oC, -15 oC, -10 oC, -5 oC,

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0 oC and 30 oC, respectively. For comparison, a chitosan hydrogel was prepared by dissolving

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chitosan powders in 2 wt% acetic acid, and then regenerating in 1 mol/L NaOH aqueous

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solution. The film was fabricated according above method, coded as CS-Na.

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To prepare the oriented CS-E films, 3 cm wide strips were clamped on a tension clamp

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and stretched at stretch rate of 0.002/s until the strain reached 10, 20, 30, or 40%,

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corresponding to draw-ratio (DR) of 1.1, 1.2, 1.3 and 1.4, respectively, and dried in air at

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ambient temperature to obtain oriented CS-E films.

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Characterization. The fracture section of the CS-E hydrogels was observed by using a

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scanning electron microscope (FESEM, Zeiss, SIGMA) at an accelerating voltage of 5 kV.

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The chitosan hydrogels were frozen in liquid nitrogen and snapped immediately, and then

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freeze-dried. The fracture surface of chitin films were sputtered with gold, and then observed 6


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and photographed. Topographic images of the diluted chitosan solution were recorded using a

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commercial atomic force microscope (AFM, Cypher ES, Asylum Research) in AC mode at

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room temperature. The optical transmittance of the CS-E and CS-Na hydrogels were observed

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with a UV-vis (UV-6, Shanghai Meipuda instrument Co., LTD., Shanghai, China)

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spectrometer at a wavelength from 200 to 800 nm. The textural properties of the chitosan

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films were determined by physically adsorbing N2 at 77 K using a physical adsorption

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analyzer (ASAP 2020M, Micromeritics Company, USA). Wide-angle X-ray diffraction

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(WAXD) patterns of the films were recorded on a Bruker APEX DUO diffractometer operated

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in reflection mode with Cu Kα radiation (λ=1.542 Å) at 40 kV and 40 mA. The crystallinity

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index (CrI) of the chitosan samples was determined using the equation30 CrI =

11

× 100%

(2)

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Where I110 is the maximum intensity (2θ=20o) of the (110) lattice diffraction and Iam is the

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intensity of amorphous diffraction at 2θ=16o of chitosan. Two-dimensional diffraction patterns

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were recorded by mounting the film sample either parallel (cross-section plane) or

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perpendicular (surface plane) with respect to the incident beam. Azimuthal intensity

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distribution from the equatorial (110) reflection in the diffractograms was extracted using

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XRD 2DScan software.

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The tensile strength (σb) and elongation at break (εb) of the films were measured on a

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universal testing machine (CMT6350, Shenzhen SANS Test Machine Co., Ltd., Shenzhen,

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China) according to ISO527-3-1995 (E) at a speed of 2 mm/min-1. The films were kept at 75%

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relative humidity for 24 h before testing. The values of mechanical properties list in table

22

represented averages of three measurements. The Young’s modulus was calculated from the 7


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initial linear region of the stress–strain curves. The experimental data were represented as

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mean ± standard deviation (SD), n=3. Statistical differences among the experimental groups

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were determined by Student’s t-test and one-way analysis of variance. Results were

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considered statistically significant when p < 0.05.

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In vitro cell assay. Cytocompatibility studies were assessed using MTT assay and carried out

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using osteoblastic MC3T3 cells, which were cultured in Minimum Essential Medium α

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(MEM α) supplemented with 10% fetal bovine serum (FBS) and 1% streptomycin/penicillin.

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The CS-E were cut into powder and sterilized by autoclaving and then used to prepare the

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extract with different concentration (0, 0.2, 0.5, 1 mg/ml) of chitosan in PBS. According to

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ISO 10993-5, osteoblastic MC3T3 cells were resuspended in the culture medium and plated

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(200 µL/well) into 96-well micrometer plates at 4 × 104 cells/mL, which were incubated at

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37 °C in a 5% atmosphere for 24 h. Then, 50 µL/well sterilized extract with different

13

concentration was added in and incubated with the culture medium itself as a control,

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eighteen replicate wells were used. After incubating for 1d, 2d, 3d, 20 µL MTT (5 mg/mL in

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PBS ﬁltered for sterilization) was added into each well and incubated for a further 4 h at

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37 °C in a 5% CO2 atmosphere, then the MTT was removed and 200 µL/well of dimethyl

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sulfoxide (DMSO) was added to dissolve the formazan crystals. The plates were placed in an

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incubator at 37 °C to shake for 10 min, subsequently, the absorbance values were read at a test

19

wavelength of 490 nm. Cell viability was calculated using the following equation:

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!"# $!"#"!% = &

&'()'

× 100%

(3)

*+,'-+.

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Where Atest and Acontrol are the absorbance values of the test and control groups, respectively.

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Data are expressed as the mean ± SD, n=18. Statistical differences between the test groups 8


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and control group were determined by Student’s t-test. The results were considered

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statistically significant when p < 0.05.

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The CS-E films (DR1.4) with small diamond shape (1cm×1cm) were sterilized by

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soaking with ethanol, then UV irradiation for 2 hours. Osteoblastic MC3T3 cells (2 × 105

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cells/well) were seeded onto the films, which were put into 24-well plates or seeded into

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plates directly as control. After incubation for 24 h, the MC3T3 cells was fixed in 4%

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formaldehyde/PBS fixative for 4 hours at room temperature, and then the cells were stained

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with Phalloidin-iFluor 647 and DAPI, to observe by using a Leica DMi8 fluorescence

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microscope. To draw the polar plots, the center of the picture was chosen as the specify origin,

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then the “Distance” is the relative distance between the cells and the specify origin, and the

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“Angle” is the angle between the long axis of the cell and the oriented growth direction, as

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shown in Scheme S1.

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RESULTS AND DISCUSSION

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Construction and structure of chitosan nanofibrous films.

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Different from the traditional chitosan materials prepared from dissolving in acetic acid

17

aqueous solution, the CS-E hydrogels and films were directly constructed by regenerating the

18

chitosan/alkali/urea solution in ethanol at low temperature. The chitosan chains in the

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LiOH/KOH/urea aqueous solution were demonstrated to easily self-aggregated to form

20

nanofibers. To provide direct evidence on the formation of the chitosan nanofibers, the diluted

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chitosan solution before and after regenerating in ethanol at low temperature were visualized

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with AFM, as shown in Figure 1. Even in an extremely dilute solution (1×10-7 g/mL), the 9


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chitosan chains and their aggregates co-existed (Figure 1a), as a result of the easy aggregation

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of the extended chitosan chains in a parallel manner in the alkaline solution.24, 30, 32 This was

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further supported by the result of DLS (Figure 1d). There were two peaks in the

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hydrodynamic radius distribution curve, corresponding to the single chitosan chains and their

5

aggregates. Interestingly, the chitosan nanofibers with mean diameter of about 20 nm were

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observed in the chitosan solution (1 ×10-5 g/mL) with cold ethanol (-10 oC) (Figure 1b).

7

Moreover, relatively larger nanofibers with mean diameter of over 33 nm appeared at the

8

chitosan concentration of 1 ×10-4 g/mL (Figure 1c). This suggested that the chitosan

9

nanofibers and bundles occurred in the concentrated solution. Thus, after regenerating in

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ethanol at low temperature the chitosan hydrogels formed, and dry films were obtained after

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air-drying (Figure 1e). The chitosan films exhibited homogenous nanofibrous and

12

hierarchically porous structure with the apparent pore size from 2 to 100 nm (Figure 1f and g).

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SEM images of the cross-section of the CS-E hydrogels are shown in Figure 2. These films

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were consisted of nanofibers with a relatively narrow pore width (Figure 2d and e). The

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average diameter of the pores was tunable from 74 to 162 nm by raising the regeneration

16

temperature from -20 to 0 oC (Figure 2f). Concretely, the lower temperature led to the smaller

17

pores and more compact structure of the films. Serious phase separation occurred at elevated

18

temperature to 30 oC, and no nanofibrous structure was observed (Figure S1). This could be

19

explained that lower temperature slowed the solvent diffusion, reduced the regeneration rate,

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so the chitosan chains could sufficiently arrange in parallel to form compact and perfect

21

nanofibrous structure, based on the Arrhenius equation.33 At the regenerating temperature of

22

-10 oC, the structure and size of nanofibers and pores were slightly influenced by the 10


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coagulant concentration (i.e. ethanol/water ratio). The average diameter of the nanofibers and

2

pores slightly decreased with increasing the concentration of ethanol (Figure S2). However,

3

nanofibrous structure disappeared and serious phase separation occurred, when the ethanol

4

concentration reduced to 50 wt% or lower (Figure S3). This revealed that the chitosan

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nanofiber easily formed with higher ethanol concentration (≥ 60 wt%). As a result, with

6

increasing the regenerating temperature or decreasing the ethanol concentration, the pore size

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of CS-E and diameter of nanofibers increased, suggesting that the lower temperature is

8

beneficial for the formation of finer nanofibrous structure.

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In view of the above results, a schematic illustration for the formation of chitosan

10

hydrogels woven with chitosan nanofibers and bundles is proposed in Scheme 1. The

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extended chitosan chains and their aggregates co-existed in the dilute solution (Scheme 1a).

12

By regenerating in ethanol, the alkaline-urea complex as shell on the chitosan was destroyed,

13

and the chitosan chains were exposed to self-aggregate in parallel to form nanofibers (Scheme

14

1b). Chitosan hydrogels could be obtained by regenerating method in aqueous ethanol (≥ 60

15

wt%) at low temperature (≤ 0 oC), in which the chitosan nanofibers and bundles aggregated

16

and entangled to form physical cross-linking networks through the intermolecular hydrogen

17

bonding (Figure 1c). The low regenerating temperature and/or high concentration of ethanol

18

were mild condition for the slow molecule diffusion to break the alkaline-urea complex shell,

19

so the chitosan chains had sufficient time for the orderly arrangement, leading to the

20

formation of the perfect nanofibers. Moreover, the chitosan hydrogels could be also

21

regenerated in methanol solution at low temperature (Figure S4). Obviously, the regenerated

22

method for the formation of the chitosan hydrogels had the certain universality. The CS-E 11


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hydrogels were drawn (Scheme 1d-f) and then used to culture cells (Scheme 1g). Thus, a

2

completely new avenue to construct the chitosan nanofibrous hydrogels and films with good

3

orientation structure from the chitosan solution in alkali/urea aqueous system was opened up

4

for the wide applications.

5

Physical and mechanical properties

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Physical and mechanical properties of materials are very important to their applications.

7

The photographs and optical transmittance curves of the CS-E and CS-Na hydrogels and films

8

are shown in Figure 3. All CS-E films exhibited much larger optical transmittance than

9

CS-Na films. This could be explained that the CS-Na films were fabricated on the basis of

10

violent neutralization, namely the severe fluctuation in the system, leading to serious phase

11

separation and inhomogeneous porous structure (Figure S5). According to the nitrogen

12

adsorption and desorption isotherms, the CS-E hydrogels showed an IUPAC type IV H3

13

hysteresis loop,34 had a specific surface area of 119.3 m2g-1 and 116.9 m2g-1 for CS-E-15 and

14

CS-E-10, respectively (Figure S6). However, CS-Na exhibited a smaller specific surface area

15

of 77.4 m2g-1. Furthermore, the Barrett-Joyner-Halendar (BJH) results indicated that the CS-E

16

possessed hierarchical porous structure with pore size in the range of 2-100 nm (Figure 1g).

17

The hierarchical porous structure could contribute to the adhesion and growth of cells,

18

transportation of nutrients, and removal of metabolic byproducts.25 XRD patterns of chitosan

19

powders and chitosan films (Figure S7) displayed only two reflection peaks, i.e. (020) and

20

(110), suggesting that the original chitosan chain packing was restructured after dissolution

21

and regeneration. The crystallinity index (CrI) was determined and listed in Table S1. The CrI

22

values (54-57%) of CS-E films were higher than that of CS-Na (44%), and increased with a 12


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decrease of regenerating temperature. The results supported the conclusion from SEM,

2

namely more regular nanofibrous structure of CS-E occurred at lower temperature.

3

Figure 4 shows the tensile stress-strain curves of the CS-E hydrogels and films. CS-E

4

hydrogels exhibited unexpectedly high tensile strength (σb=3.76 MPa), elongation at break

5

(εb=45.8%), and Young’s modulus (E=5.15 MPa), significantly higher than that of CS-Na

6

(0.08 MPa, 14.0%, 0.46 MPa) (Figure S8). Moreover, the CS-E films exhibited the highest σb,

7

εb and E values (144.8 MPa, 28.3%, 3.5GPa) among the as-reported pristine chitosan films

8

(Table S1).35-37 The mechanical properties of the CS-E hydrogels and dry films were strongly

9

influenced by the regenerating temperature. The decreasing of the regenerated temperature

10

could significantly enhance the tensile strength, modulus of elasticity and elongation at break

11

decreased (Figures S9 and S10, Table S1), further supporting that low temperature was

12

beneficial to the sufficient order-arrangement of the chitosan chains. Moreover, the CS-E

13

prepared at lower temperature had higher σb and εb values, consistent with the results of SEM

14

and XRD (Figure 2 and S7). The CS-E films not only had high tensile strength but also

15

displayed remarkable toughness, as a result of the unique nanofibrous structure. The excellent

16

mechanical properties of the chitosan films were as a result of the strong networks woven

17

with the compact nanofibers. Interestingly, the CS-E films could be bent and folded arbitrarily

18

to various complex shapes, and these shapes could be retained for a long time, like cellulose

19

paper. As shown in Figure 3, the CS-E hydrogels were knotted as a necktie, and CS-E dry

20

films were folded as a crane, which guaranteed the application of CS-E films. In contrast,

21

CS-Na films ruptured easily with folding (Figure S11), revealing a weak and fragile feature,

22

as a result of the inhomogeneous structure. The good mechanical properties of the CS-E films 13


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and hydrogels were important in the biomedical applications.

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Orientation by drawing

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The mechanical strength of polymer films could be improved greatly by drawing.

4

Usually, the uniaxial drawing leads to a significant reorientation of the polymer chains in line,

5

resulting in an enhancement of the strength.38 The CS-E films with oriented nanofibrous

6

structure were prepared by cold-drawing herein. As shown in Figure 5a, the CS-E hydrogels

7

without drawing (RT1.0) have a random or non-oriented nanofiber structure, and showed an

8

oriented drawn structure after drawing. In Figure 5b, the dried CS-E films displayed typical

9

ring patterns of the chitosan crystals with two reflections corresponding to (020) and (110) in

10

the surface plane (SP), consistent with the result of Figure S6, indicating the random

11

distribution on the nanofibers. In contrast, the CS-E film showed equatorial arcs

12

corresponding to (020) and (110) in the cross-sectional plane (CP). This could be explained

13

that the nanofibers aligned in parallel on the film surface. However, the two reflections of

14

oriented films with draw ratio of 1.4 displayed equatorial arcs in the SP, revealing the

15

alignment of chitosan nanofibers in the SP along the drawing direction. Meanwhile, the CS-E

16

film after drawing exhibited a pattern with colors when viewed between the cross polarizers,

17

different from that of before drawing (Figure S12), suggesting the anisotropic structure of the

18

drawn CS-E film.

19

The (110) reflection was used to quantify the chitosan crystal orientation, and can also be

20

used to evaluate the orientation of the nanofibers. The orientation index (fc) of the crystals

21

was calculated to quantify the nanofiber orientation in the films by using azimuthal breadth

22

analysis, according to the following equation39 14


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f0 = (180 − 4567)/180

1

(4)

2

Where FWHM is the half-width of the azimuthal distribution curve along the equatorial (110)

3

reflection in the X-ray diffractograms. The azimuthal distribution curves of (110) plane of the

4

films are plotted in Figure 5c, and the orientation index are listed in Table S2. The fc value in

5

both SP and CP directions increased with the increasing of the drawing ratio. The oriented

6

CS-E film (DR1.4) had the fc value of 83.9 and 84.0 for SP and CP directions, respectively,

7

even higher than those of the wet-spun fibers from wood cellulose nanofibers (fc between 0.65

8

and 0.72).40 In addition, a 532 nm single mode laser was used as the incoming light source for

9

the anisotropy measurement.39 As shown in Figure 5d, laser beam travels through CS-E

10

without drawing and obtained a small and intense spot on the wall, and the light scattering

11

was negligible. After drawing, the illuminated area increased and homogenous light scattering

12

pattern appeared. Interestingly, the scattering was biaxial and resulted in an elliptic shape of

13

illuminated area on the wall with a large draw ratio, as a result of the alignment of chitosan

14

crystal in the films along the drawing direction. In view of the results, the resulted CS-E

15

hydrogel films could be easily oriented by drawing to achieve a maximum orientation index

16

of 84 %.

17

Tensile stress-strain curves and mechanical properties of oriented CS-Es are presented in

18

Figure 6a and Table S2, respectively. With an increase of draw ratio, the tensile strength and

19

modulus of elasticity of the oriented CS-E films significantly increased, whereas the

20

elongation at break reduced (Figure S13). As shown in Figure 7b, the tensile strength and

21

elongation at break of the oriented CS-E were 282 MPa and 9.3 %, respectively, which were

22

the highest values among the reported chitosan films.36, 41-43 This could be explained that the 15


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

CS-E films composed of nanofibers had homogeneous architecture and strong networks

2

weaved with nanofibers. In our finding, the cold-drawing process further improved the

3

mechanical properties of the CS-E films. Therefore, a novel approach to construct the oriented

4

chitosan nanofibrous films with excellent strength via a “bottom-up” pathway was

5

established.

6

Growth of cell on oriented chitosan hydrogel films

7

Biomaterials with oriented structure are usually necessary in biomedicine or tissue

8

engineering for the directional growth and migration of cells. For example, aligned fiber films

9

have been used to relocate or manage the growth of primary tumors to achieve the purpose of

10

treatment.21 The in vitro cytotoxicity tests on the CS-E films (Figure S14) showed that the cell

11

viability values on all chitosan films were higher than 96%, indicating no cytotoxicity. Further,

12

the oriented CS-E films as tissue engineering scaffolds for cell adhesion were evaluated.

13

Usually, cells are in micro-scale, whereas the orientation of CSE was in nanoscale. However,

14

in our findings, the adaptation and growth of the discrete filopodia of the cells along the

15

oriented nanofiber direction (Figure S15). Moreover, it has been demonstrated that the L02

16

cells filopodium can extend and adhere on the nanofibers of the peripheral microspheres.25

17

Thus, osteoblastic MC3T3 cells were seeded on test hydrogels for 24 hrs to assess the

18

morphology and alignment, because there are too many cells to observe obviously directional

19

growth if incubated for longer time. And the CS-E without orientation was used as control.

20

Morphologies of cells on the different substrates were analyzed by applying cytoskeleton and

21

nucleus staining with phalloidin and DAPI, respectively, with fluorescence microscopy

22

imaging (Figure 7). Obviously, MC3T3 cells could adhere, spread, and proliferate well on the 16


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1

surface of the hydrogel films. The osteoblasts grown on both CS-E and oriented CS-E

2

exhibited elongated shapes, similar to the cells on the petri dish substrate. Interestingly, the

3

cells on the oriented CS-E substrate grew directionally, which could be correlated with the

4

orientation direction. The angle distribution (Figure 7b) indicated that the orientation of the

5

cells on CS-E without orientation was same as that on the petri dish substrates, which were

6

completely random and covered all possible angles. However, the cells distributed on the

7

oriented CS-E substrate were concentrated in the angle range of -30o to 30o, on the whole.

8

This demonstrated that the oriented film could induce the directional growth of both normal

9

cells and tumor cells (Figure 7a, Scheme 1g and Figure S15). About 78% of the cells on

10

oriented CS-E substrate were aligned within 20o from the orientation directions (Figure 7c),

11

which were higher than that of the reported aligned carbon nanotubes (70% of cells aligned

12

within 20o).44 In view of these results, the growth direction of the cells could be regulated by

13

the oriented structure of CS-E, on the basis of the discrete filopodia of the cells to extend and

14

adhere on the nanofibers.

15 16

CONCLUSION

17

A facile route to fabricate chitosan nanofibrous hydrogels and films from chitosan

18

solution in alkali/urea aqueous system via a mild regeneration process was established

19

successfully. The high strength nanofibrous hydrogels and films could be formed at low

20

regenerating temperature (≤ 0 oC) and relatively high ethanol concentration (≥ 60wt%), in

21

which the extended chitosan chains easily self-aggregated in parallel to form the relatively

22

prefect nanofibers, as a result of the sufficient order-arrangement of the chitosan chains in the 17


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1

mild system. Thus, the nanofibrous chitosan films exhibited high strength, and could be fold

2

to various complex shapes. Via simply drawing, the oriented CS-E films with ultra-high

3

tensile strength and modulus were fabricated, showing high degree of nanofibrous orientation.

4

Moreover, the CS-E hydrogels possessed the hierarchically porous structure, which was

5

important in the application of tissue engineering. The cell viability values on all chitosan

6

hydrogels were higher than 96%, indicating no cytotoxicity. The osteoblastic cells could

7

adhere, spread, and proliferated well on the CS-E hydrogels, indicating good biocompatibility.

8

Furthermore, the oriented CS-E could induce the growth of cell in a certain direction, on the

9

basis of the filopodia of the cells to extend and adhere on the nanofibers.

10 11

ASSOCIATED CONTENT

12

SUPPORTING INFORMATION

13

Graphical illustrationfor the drawing of the polar plot; SEM images of the cross-sectional

14

structures of CS-Es regenerated in different ethanol solution; nitrogen adsorption-desorption

15

isotherm of CS-E and CS-Na hydrogel films; XRD patterns of chitosan powders, CS-E and

16

CS-Na films; Pictures of CS-Na hydrogels and films after folding; the dependence of relative

17

activity of cells on the concentration of chitosan films; physical and mechanical properties of

18

the chitosan films. This material is available free ofcharge via the internet at

19

http://pubs.acs.org.

20 21

ACKNOWLEDGEMENTS

22

This work was supported by the Major Program of National Natural Science Foundation of 18


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Biomacromolecules

1

China (21334005), the Major International (Regional) Joint Research Project (21620102004)

2

and the National Natural Science Foundation of China (51573143, 51203122).

3 4

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Chem. 2013, 15, 3404-3413.

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2017, 5, 2725-2733.

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Hydrocolloids 2016, 61, 662-671.

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1

Biomacromolecules

FIGURE CAPTIONS

2 3

Scheme 1. Graphical illustration for the formation of the nanofibers and networks of the

4

chitosan hydrogel films (CS-E) from their solution: coexistence of the CS-alkaline-urea

5

complexes and their aggregates in the solution (a); the formation of the perfect chitosan

6

nanofibers in ethanol solution (≥ 60 wt%) at low temperature (≤ 0 oC) (b); the formation of

7

the physical cross-linking networks consisted of the CS nanofibers and bundles (c). Schematic

8

illustration for the preparation process of the oriented CS hydrogels and directional growth of

9

cells (d). Picture of the drawing CS-E hydrogel (e). SEM images of the surface of the oriented

10

CS-E hydrogel (f) and the rat glial tumor (C-6) cells spreading on the oriented CS-E hydrogel

11

(g).

23


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2

Figure 1. AFM images of the extremely dilute chitosan solution in alkaline system

3

(1×10-7 g/ml) (a), and chitosan aggregates regenerated in ethanol from the chitosan

4

solution with a concentration of 1×10-5 g/ml (b) and that with 1×10-4 g/ml (c) at -10

5

o

6

(8×10-4 g/ml) at 5 oC (d). Picture of the CS-E film (e), SEM image of the

7

cross-section (f) and Barrett-Joyner-Halendar (BJH) pore size distribution (g).

C. The hydrodynamic radius distributions [f(Rh)] of the chitosan alkaline solution

8 9 10 11 12 13 14 15 16 24


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1 2

Figure 2. SEM images of the CS-E hydrogel films prepared in 80 wt% ethanol

3

solution at different regeneration temperature: -20 oC (a), -10 oC (b), 0 oC (c). Pore

4

width distribution of CS-E-20 (d) and CS-E0 (e). Pore width of CS-Es regenerated at

5

different temperature (f).

6 7 8 9 10 11 12 13 14 15 16 17 18 25


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2

Figure 3. The pictures of the CS-E hydrogels (a, b) and films (c), as well as the

3

CS-Na hydrogels (d), and the optical transmittance curves of the CS-Es and CS-Na

4

hydrogels in the range of visible light (f).

5 6 7 8 9 10 11 12 13 26


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1 2

Figure 4. Mechanical properties of the CS-E, CS-Na hydrogels (a) and films (b).

3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 27


Biomacromolecules

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1 2

Figure 5. SEM images of the surface of the CS-E hydrogel before and after drawing

3

(bar: 500 nm) (a), X-ray diffractograms of the SP (beam perpendicular to the surface)

4

and CP (beam parallel to the surface) direction of the CS-E films with different draw

5

ratio (b), azimuthal intensity distribution profiles of the (110) scattering plane

6

obtained from X-ray diagrams of the films in the SP direction (c), and light scattering

7

images of the CS-Es with different draw ratio (d). The diameter of the laser point is 1

8

mm, the distance between the film and the wall is 30 cm.

9 10 11 12 13 28


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1 2

Figure 6. Effects of draw ratio on mechanical properties of the CS-E films (a) and

3

comparison of mechanical properties of various chitosan films (b) 36, 41-43.

4 5 6 7 8 9 10 11 12 13 14 15 16

29


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2

Figure 7. The adhesion and growth of the osteoblastic MC3T3 cells on the chitosan

3

hydrogels: fluorescence microscopy images of the cells cultured on petri dish (top),

4

CS-E (middle) and oriented CS-E (bottom) stained for F-actin (left, red) and nucleus

5

(right, blue), scale bar: 100 µm (a), polar plot of the orientation of cells on the petri

6

dish (black square), CS-E (red circular) and oriented CS-E (blue triangle). In the polar

7

plot, the “Distance” is the relative distance between the cells and the specify origin,

8

and the “Angle” is the angle between the long axis of the cell and the oriented growth

9

direction (b). Angle distribution (no distinction of positive and negative) histograms

10

of the cells cultured on different substrates (c).

11

.

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1 2

Biomacromolecules

High-Strength Films Consisted of Oriented Chitosan Nanofibers for Guiding Cell Growth

3 4

Kunkun Zhu, Jiangjiang Duan, Jinhua Guo, Shuangquan Wu, Ang Lu*, Lina Zhang*

5 6

College of Chemistry and Molecular Sciences, Wuhan University, Wuhan 430072, China

7

*Correspondence to: [email protected] (L. Zhang), [email protected] (A. Lu)

8

9 10 11 12

The high strength nanofibrous hydrogels and films were fabricated from the chitosan solution

13

in LiOH/KOH/urea aqueous system via a mild regenerating process, in which the extended chitosan

14

chains sufficiently self-aggregated in parallel to form the prefect nanofibers. The CS-E hydrogels

15

could be easily oriented by drawing, and the oriented chitosan hydrogel films could efficiently

16

regulate the directional growth of osteoblastic cells along the orientation direction.

17 18

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High-Strength Films Consisted of Oriented Chitosan Nanofibers for

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