Mechanically Strong Chitin Fibers with Nanofibril Structure

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Mechanically Strong Chitin Fibers with Nanofibril Structure, Biocompatibility and Biodegradability Kunkun Zhu, Hu Tu, Pengcheng Yang, Cuibo Qiu, Donghui Zhang, Ang Lu, Longbo Luo, Feng Chen, Xiangyang Liu, Lingyun Chen, Qiang Fu, and Lina Zhang Chem. Mater., Just Accepted Manuscript • DOI: 10.1021/acs.chemmater.8b05183 • Publication Date (Web): 25 Feb 2019 Downloaded from http://pubs.acs.org on February 26, 2019

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Chemistry of Materials

Mechanically Strong Chitin Fibers with Nanofibril Structure, Biocompatibility and Biodegradability Kunkun Zhu1#, Hu Tu1#, Pengcheng Yang3, Cuibo Qiu2, Donghui Zhang3, Ang Lu1*, Longbo Luo2, Feng Chen2, Xiangyang Liu2, Lingyun Chen4, Qiang Fu2*, Lina Zhang1*

1College

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

2College

of Polymer Science and Engineering, Sichuan University, Chengdu 610065, China

3State

Key Laboratory of Biocatalysis and Enzyme Engineering, School of Life Science, Hubei

University, Wuhan, Hubei 430062, China 4Department of Agricultural, Food & Nutritional Science, University of Alberta, Edmonton, AB, T6G

2P5 Canada

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

ABSTRACT Facing the global pollution of the fibers and textile fabricated from petroleum-based polymers, the environmental friendly fibers fabricated from natural polysaccharides have attracted much attentions for the development of the sustainable materials. Chitin derived from seafood wastes possesses excellent biocompatibility and biodegradability, but it is still far from fully explored. Here, we designed and prepared, for the first time, the chitin fibers with nanofibril structure from the chitin solution in the NaOH/urea aqueous system with cooling on a lab-scale wet-spinning machine. Due to the slow diffusion of phytic acid into the chitin dope, the stiff chitin chains could self-aggregate 1

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sufficiently in parallel to form the nanofibers via a “bottom-up” approach, and then bundled into the gel-state fibers. The dried chitin fibers were demonstrated to be consisted of nanofibers with mean diameter of 27 nm, and exhibited a tensile strength of 2.33 cN/dtex, higher than those reported in literatures. Interestingly, with an increase of the draw ratio from 1.0 to 1.8, the crystal index (χc) and degree of orientation (Π) increased very slightly, whereas the tensile strength and Young’s modulus of the chitin fibers enhanced significantly, suggesting that relatively perfect nanofibrous structure existed in all of chitin fibers with and without drawing. Moreover, the chitin fibers were validated to support the adhesion and growth of ventricular myocytes as cardiac tissue scaffold, showing good biocompatibility. Furthermore, the complete biodegradation time of the chitin fibers in soil and in vitro could be extrapolated from experimental data to be approximately 22 and 34 days, respectively, indicating good biodegradability. This work would lead to a great potential of chitin in the applications including absorbable surgical suture, hemostasis and fixation medical device etc., where biodegradability is required.

INTRODUCTION It is well- known that coal and oil will be running out in future, and the petroleum based polymer materials in the form of fiber, films, sponges, plastics have caused great pollution to the environment due to their difficult biodegradations. Particularly, the oceans are polluted from the plastic wastes, which are considered as hazardous,1-3 and if the current trend continues, there could be more plastic than fish in the oceans by 2050.4 Most of the natural polysaccharides such as chitin, alginate and cellulose derived from the ocean biomass, from which sustainable materials are generated to show nontoxicity, biocompatibility, biodegradability, and high tensile strengths.5-11 These polysaccharides based materials are appropriate for the biomedical applications, specifically, chitin extracted mainly 2


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from crab and shrimp wastes is abundant, and has significant potential in biomaterials as an original component of living organisms.12, 13 Faced on a great technical challenge to endow biomaterials with excellent properties, new biocompatible, biodegradable, and tough polysaccharides based biomaterials have attracted much attentions.14-16 In the shell of crab and shrimp, their fibers are consisted of chitin nanofiber wrapped in protein layers.17 Meanwhile, cost effective and environmentally benign approach to separate the chitin from the shell is viable using only water and CO2, beyond the conventional NaOH/HCl process.18 The chitin nanofibers play important role as the basic structural unit for the toughness of the naturally occurring materials, and so inspired us to fabricate the nanofibril structure via a bottom-up approach from its solution. Bottom-up approach is a method that built smaller (usually molecular) components up into more complex assemblies. However, chitin is hardly soluble and difficult to be processed, which made it less exploited.19 To date, only a few solvents such as dimethylacetamide (DMAc)-LiCl mixtures,20 hexafluoro isopropanol (HFIP),21, 22 CaCl2-MeOH23, 24 and ionic liquids25, 26 have been acceptable to dissolve chitin. In our laboratory, NaOH/urea aqueous solution, an environmentally friendly water based solvent, has been developed to dissolve chitin at low temperature, based on the formation of chitin-alkali-urea complexes through the forming hydrogen bonding between macromolecules and solvents. In this system, the chitin exists as a sheath-like structure adopting an extended chain conformation, and the single chitin molecules and their aggregates co-exist in the aqueous solution.27, 28 Moreover, a series of nanofibril-structured hydrogels, films, microspheres and aerogels have been directly constructed from the chitin solution.29-32 However, the chitin fibers with nanofibril structure have been never reported. In this work, chitin fibers were prepared by wet-spinning on a lab-scale spinning machine. Phytic acid was chosen as a coagulant to mitigate the regeneration process due to its low self-diffusion 3



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coefficients, which plays an important role in the formation of nanofibril-structure.33 The hydronium ions in phytic acid can relatively slowly diffuse into the chitin solution to destroy the alkali-urea hydrogen bonded complex shell, and then the naked stiff chitin chains can self-aggregate sufficiently in a parallel manner (with the largest contact area) to form the nanofibers via a “bottom-up” approach.9, 28, 33 The structure of the chitin fibers was characterized by 13C NMR, scanning electron microscopy (SEM), X-ray diffraction (XRD), two-dimensional wide-angle X-ray diffraction (2D WAXD) and small angle X-ray scattering (SAXS). The mechanical properties and the biodegradability of the chitin fibers were evaluated by tensile testing and the degradation in soil and in vitro. Moreover, the cell culture test was conducted to evaluate the potential of the fibers for biomaterial applications. This work is important to fabricate highly strong chitin fibers with biocompatibility and biodegradability, and assessed the possibility of replacing the not biodegradable polymer fibers. For example, it may be used as bioabsorbable materials and medical devices which can be biodegraded in the biological environment in vivo and under natural soil, to the benefit of both avoiding secondary surgery in in clinical use and protecting environment.

EXPERIMENTAL SECTION Materials. Commercial chitin powder was purchased from Golden-Shell Biochemical Co., Ltd. (Zhejiang, China), and was purified according to a previously reported method.34 The weight-average molecular weight (Mw) was measured by light-scattering spectrometer (ALV/SP-125, Germany) in NaOH/urea aqueous system at 20 oC to be 3.2 ×105 g/mol. All other chemical reagents were purchased from Sinopharm Chemical Reagent Co., China and used without purifications. Preparation of Chitin Fibers. To prepare the spinning dope, chitin powder (120 g) was added into the alkaline aqueous solvent (1880 g) containing NaOH/urea/H2O of 11:4:85 by weight. The resultant 4


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suspension was frozen at -30 oC for 4 h, and then thawed at room temperature. The freezing-thawing cycle was repeated twice to dissolve the chitin thoroughly. Then 6 wt% chitin dope was prepared after centrifugation at 5 oC. The wet-spinning process was performed at room temperature (30 oC) on a lab-scale wet-spinning machine (Figure S1) according to the early report,33 in which a spinneret with 50 orifices and diameter of 160 μm was used. The chitin dope was pushed into a coagulation bath of 15 wt% phytic acid/5 wt% sodium sulfate aqueous solution through spinneret from a sealed reservoir into a Zenith BPB-4391 gear pump using nitrogen pressure (0.15 MPa). The gel-state fibers regenerated in the first coagulation bath were taken up on the Nelson-type roller I, and then drawn to the Nelson-type roller II in the second coagulation bath, which is hot water. Then the gel-state fibers were washed to remove the residual salts and acids. After drying on the heating roll (60 oC), the drystate chitin fibers were collected on the take-up roller. The flow rate of the chitin solution through the spinneret holes was 7.46 m/min. The multi-drawing processes were achieved through Nelson-type rollers I and II and take up roller. The code of the fibers fabricated with different wet spinning parameters are summarized in Table S1. The Characterization. SEM images were taken on a scanning electron microscope (FESEM, Zeiss, SIGMA) at an accelerating voltage of 5 kV. The gel-state chitin fibers were snapped immediately after frozen in liquid nitrogen, freeze-dried, and then sputtered with gold to observe the cross-section. The inner pattern of the chitin fibers was firstly embedded in epoxy resin and then sliced along the fiber axis by microtome to prepare a slice sample (10 μm thick), which was sputtered with gold before observe. The cross-section of the chitin fibers was observed on an optical microscope (Leica DMLP, Germany). Solid-state 13C NMR spectra were examined on a BRUKER ACANCE III spectrometer operated at a

13C

frequency of 75 MHz using the combined technique of magic angle spinning (MAS) and 5



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cross-polarization. The spinning speed was set at 5 kHz. The contact, acquisition and delay times were 3 ms, 50 ms and 3 s, respectively. A typical number of 1024 scans were acquired for each spectrum. The chitin fibers were cut into powders and dried before measurement. XRD of the chitin fibers were recorded with a Rigaku Miniflex600 diffractometer in reflection mode with Cu Kα radiation (λ= 0.154 nm). The scanning speed was 5 oC/min and the 2θ ranged from 4o to 40o with a step-size of 0.02o. The chitin fibers were cut into powders to eliminate the effects of the crystalline orientation. The crystallinity of the samples was calculated by using the following equation35 𝐴𝑐𝑟

𝜒𝑐(%) = 𝐴𝑐𝑟 + 𝐴𝑎𝑚 × 100

(1)

Where Acr and Aam are the integrated area of the crystalline and amorphous phases, respectively. 2D WAXD measurements were carried out on a Bruker D8 Advance diffractometer with a Cu anode via the Debye-Scherrer method. The generator was operated at 40 kV and 0.65 mA. The degree of orientation (Π) was calculated by using the following equations36 𝛱=

180 ― 𝑓𝑤ℎ𝑚

(2)

180

Where fwhm is the full width at half-maximum of the azimuthal distribution curve along the equatorial (020) reflection. SAXS patterns were obtained on a MP-Xeuss 2.0 SAXS (BRUKER AXS, Inc.) operated at 40 kV and 0.65 mA with Cu Kα radiation (λ= 0.154 nm) by using a HI-STAR detector and the specimen to detector distances were 1074 mm. The chitin fibers were straightened to bundle before the measurement. The linear density of the chitin fiber was calculated in terms of dtex, which are defined as the weight (g) per 10,000 m. Mechanical properties of the chitin fibers were tested on a universal tensile tester (LLY-06ED, Laizhou Electronic Instrument Co., Ltd. China) at 23°C and humidity of 65% 6


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according to ISO527−3−1995 (E). The gauge length is 20 mm with a stretching velocity of 20 mm/min. The tensile strength (σ) and elongation at break (ε) values were obtained from at least 20 independent specimens for each fiber prototype. Neonatal Rat Ventricular Myocytes (NRVM) Isolation and Culture. Ventricles were extracted from 1-2-day-old Sprague-Dawley rats and cardiomyocytes were isolated using Neomyts kit (Cellutron, NC-6031). The sterilized chitin fibers were cut into 12 mm fragments and fixed on polydimethylsiloxane (PDMS) grooves (8mm×8mm×2mm) by nylon frames and pins (Figure S5). Hydrogel solution (24 μL human fibrinogen (10 mg/mL, Sigma F4883), 12 μL Matrigel, 32 μL Dulbecco’s modified Eagle medium (DMEM) were mixed with 1.5 × 106 cells in 48μL NRVM culture medium. Following addition of 2 μL thrombin (50 U/mL, Sigma T7513), cell/hydrogel solution was added to the grooves and left at 37 °C for 1 h to polymerize. The cell/hydrogel with chitin fibers were cultured in medium consisting of Dulbecco’s modified Eagle medium (DMEM) with 4.5 g/L glucose, 5% certified fetal bovine serum (FBS), 1% streptomycin/penicillin and 50 μg/μL ascorbic acid. After being cultured for 4 days, Cell/hydrogel with chitin were removed from PDMS grooves and cultured in 24-well plates for additional 3 days in 500 μL of cultured medium, which contained 1mg/mL aminocaproic acid to prevent fibrin degradation. The cells were stained by Sarcomeric, α-actinin (Sigma, A7811) and Dapi. Confocal images were taken by Zeiss LSM710. Biodegradation Tests. The chitin fibers enclosed in a nylon fabric (500 meshes) were buried about 10 cm depth in the natural soil, and the environment temperature was in the range from 25 to 30 oC during biodegradation test. The samples were removed one by one, cleaned carefully with water, and then dried in oven. The degraded chitin fibers from the burying time of 2 to 17 days were characterized with digital photo, SEM and the mass change from the degradation kinetics, respectively. The 3-day intervals were chosen to investigate the weight loss of the chitin fibers. 7



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In vitro degradation of chitin multifilaments was examined by gravimetric method. A certain amount chitin fibers was wrapped in nylon cloth. Then the samples were incubated in 30 mL of 0.15 M phosphate-buffered solution (PBS, pH 7.4) at 37 oC in the presence of 1.5 μg/ml lysozyme (Solarbio, Beijing, China) with the activity of 20,000 U/mg.37 Briefly, Samples were removed from PBS solution and weighed after rinsing with distilled water and drying under vacuum at several day intervals. The degradation dynamics and surface morphology of fibers were observed, and utilized to analyze the degradation activity.

RESULTS AND DISCUSSION Formation of the Chitin Nanofibers As mentioned above, the crab shell is consisted of chitin nanofibers, proteins and minerals,

17

suggesting that the chitin nanofibers may be generated directly by removing the other components via a “top-down” approach.38 Correspondingly, “bottom-up” is the other method for the nanofiber preparation, which bundles the macromolecules in solution into nanofibers.39 Herein, a simple and effective strategy to construct chitin fibers with nanofibril structure directly from the chitin dope in NaOH/urea aqueous solution by wet-spinning was proposed. Figure 1 shows the fabrication process and formation mechanism of the chitin fibers with nanofibril structure. The chitin powder was dissolved in NaOH/urea aqueous system with cooling to produce a transparent chitin solution, in which the rigid chitin molecule chains and their aggregates as nanofibers co-existed.27, 28 The wetspinning process of the chitin dope was performed on a lab-scale wet-spinning machine (Figure S1). Based on the relatively slow diffusion of phytic acid into the spinning dope spouted from the spinneret, the stiff chitin chains could self-aggregate sufficiently in parallel to form the nanofibers through strong intermolecular hydrogen bonding interaction of the chitin, and then bundled into the gel-state 8


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fibers.33, 40 In phytic acid, the NaOH-urea hydrogen bonded shell surrounding the chitin chain was destroyed, and the naked chitin chains self-aggregated easily in a parallel manner, leading to the chitin nanofiber formation via a “bottom-up” approach. After washing and drying, the colorless chitin fibers with nanofibril structure (CTF) were obtained. In our laboratory, the cellulose fibers with nanofibrilstructure have been also spun successfully through this technology.33 The multi-drawing processes were achieved by controlling the rotation rate of Nelson-type rollers I and II. According to the draw ratio of 1.0, 1.4 and 1.8, the resulted chitin fibers were coded as CTF-1.0, CTF-1.4 and CTF-1.8, respectively. Other chitin fibers with different coagulation temperatures were numbered sequentially from CF-1 to CF-5, summarized in Table S1.

Figure 1. Preparation and formation mechanism of the chitin fibers derived from crab and shrimp shells. Commercial chitin powder derived from crab and shrimps was purchased from Golden-Shell Biochemical Co., Ltd. (Zhejiang, China). After purification, the chitin powders were dissolved in NaOH/urea aqueous system with cooling to obtain the chitin solution, from which the chitin multifilament fibers were fabricated by wet spinning. 9



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Structure of the Chitin Fibers and Drawing Effects According to the optical microscopy image in Figure 2a, the cross-section of the chitin fiber was circular, similar to the cellulose fiber fabricated through the same method.33 The chitin fiber exhibited homogeneous nanofibrous and hierarchically structure (Figure 2b and c). The effect of coagulating temperature on the formation of chitin nanofibers indicated that the average diameter of nanofibers decreased from 35 to 28 nm, with decline of the coagulating temperature from 32 to 7 oC of the first coagulation bath (Figure S2, from CF-1 to CF-3). This could be explained that lower temperature led to relatively slower molecular diffusion and regeneration rate, thus the chitin chains had sufficient time to arrange in parallel to form relatively perfect nanofibers.41 The temperature of the second coagulation bath from 60 to 20 oC had no obvious influence on the nanofibril structure (Figure S2, from CF-3 to CF-5), as a result of that the nanofiber formation was completed in the first coagulation bath. Interestingly, chitin fibers regenerated in H2SO4 coagulation showed a porous structure, because of the fast regenerating process of chitin solution in H2SO4 coagulation.33, 42

Figure 2. Microscopic morphology of the chitin fibers. (a) Optical microscope images of the cross10


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section of the chitin fibers in the dry state. (b) SEM images of the cross-section of the chitin fibers in the wet state and (c) its enlarged view. (d-f) SEM images of the inner structure and (g-i) diameter distribution of the single nanofiber of (d, g) CTF-1.0, (e, h) CTF-1.4 and (f, i) CTF-1.8 in the dry state, which were prepared by slicing along the fiber axis direction.

The nanofibril structure of the chitin fibers in the wet and dry states were further validated with SEM. The nanofibers in the chitin fibers in the wet state displayed an average width of 30 nm, 300800 nm in length and 18 in average aspect ratio (Figure 2 and S2). The dried chitin fiber was epoxyresin-embedded, sliced along the fiber axis by microtome, and then observed by SEM (Figure 2d-f). Clearly, the nanofibers arranged in a parallel pattern and the broken heads decreased with the drawing process. Interestingly, the nanofibers in the chitin fiber was not individual nanofiber, and they selfaggregated to form dense aligned architecture, similar to that existed in natural woods.8 This could be explained that the chitin nanofibers easily to self-aggregate together through strong intermolecular hydrogen bonding interaction at the gel state. The nanofibers in the chitin fibers in the dry state had an average width of 27 nm (Figure 2d-i), smaller than that at the wet state because of water volatilization of the fiber during dry process. Meanwhile, little change was observed for the average diameter of the chitin nanofibers with the drawing progress as shown in Figure 2g-i. This indicated that the relatively perfect chitin nanofibers formed and fixed in the first coagulation bath, and the structure became denser under the drawing condition. To further clarify the effects of drawing, three kinds of fibers with different draw ratios, were observed with SEM, 2D WAXD and SAXS (Figure 3). With the progress of drawing, the mean apparent diameter of the chitin fibers in the wet state decreased from 94 to 69 μm and the surface become smooth (Figure 3a-c), which was also observed in the dry state (Figure S3). All the 2D 11



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WAXD patterns of the chitin fibers (Figure 3d-f) exhibited several Debye rings with narrow arcs from the outside to the center, indicating that the chitin chains were preferentially oriented with their crystallographic axis in the drawing direction.43 From SAXS patterns of the chitin fibers (Figure 3gi), long equatorial streaks and short meridional peaks appeared, suggesting the presence of needleshaped voids or fibril structure parallel aligned to the fiber direction with a periodic lamellar arrangement of the crystalline and amorphous chitin regions. The orientation degree of the chitin fibers increased very slightly from 0.77 to 0.78, corresponding to the draw ratio from 1.0 to 1.8 (Table S2).

Figure 3. (a-c) SEM images of the chitin fibers in the wet state, (d-f) 2D WAXD and (g-i) SAXS patterns of (a, d, g) CTF-1.0, (b, e, h) CTF-1.4 and (c, f, i) CTF-1.8.

13C

NMR spectra of the chitin fibers are shown in Figure 4a. The spectrum of the chitin powder

displayed eight peaks: the sharp peaks at 174.5 and 23.0 ppm were attributed to carbonyl and methyl 12


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carbons, respectively; the peaks at 104.5, 83.1, 76.0, 73.4, 60.9 and 55.1 ppm were ascribable to the resonances of C1, C4, C5, C3, C6 and C2 on the N-acetyl-D-glucosamine unit of chitin, respectively (Figure S4).44 The peaks assigned to C3 and C5 in both the chitin powder and the fibers existed as two peaks, suggesting the same α-chitin structure.45, 46 The shoulder peak at 173.3 ppm for the C7 resonance in the chitin powder disappeared in the chitin fibers, revealing the breaking of the intermolecular hydrogen bonds (C(6)-OH●●●O=C).47 However, the line width in the chitin fibers increased with the drawing process, showing stronger hydrogen bonding interactions between the chitin chains and bundles, as well as more ordered structures after drawing.48 Meanwhile, the degree of acetylation (DA) of the chitin powder and chitin fibers was calculated from the 13C NMR spectra49 to be 94% and 91-92%, respectively (Table S2), suggesting only a slight deacetylation occurred during the dissolution and regeneration process. Moreover, the chemical shifts of the eight carbon atoms of chitin fibers were similar to those of the chitin powder. The results further demonstrated that the chitin dissolution and regeneration were physical processes,50, 51 no chemical reaction occurred. This feature was also supported by the XRD spectra of the chitin powder and fibers (Figure 4b). All the chitin fibers and chitin powder exhibited six diffraction peaks at 2θ = 9.4°, 12.9°, 19.3°, 20.8°, 23.5°, 26.5° indexed as (020), (021), (110), (120), (130) and (013), respectively, suggesting the crystalline structure of α-chitin.52, 53 The diffraction peaks of the chitin fibers were much broader and weaker than that of the chitin powder, indicating a decrease in the crystallinity (Table S2). Meanwhile, the crystal index (χc) increased slightly from 74.2 to 76.8% with an increase of the draw ratio from 1.0 to 1.8. In view of these results, the chitin fibers retained the intrinsic structure and character of native α-chitin, namely native bioactivity.

13



Figure 4. (a) CP/MAS

13C

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NMR and (b) XRD spectra of purified chitin powder, CTF-1.0, CTF-1.4

and CTF-1.8. (c) Photograph of the chitin fibers and (d) stress-strain (σ-ε) curves of the chitin fibers.

The mechanical properties of materials are crucial to their wide applications. The mechanical properties of the chitin fibers with different wet spinning parameters (Table S2 and Figure S5) showed that the tensile strength increased with a decrease in temperature of the first coagulation bath (from CF-1 to CF-3), suggesting that the perfect nanofibril structure could form at relatively low temperature. However, elevating the temperature of the second coagulation bath led the stronger fibers (from CF-5 to CF-3), as a result of the quick fixing of the oriented chitin chains and nanofibers during the drawing process. With an increase of the draw ratio from 1.0 to 1.8, the tensile strength and Young’s modulus enhanced from 1.66 to 2.33 cN/dtex and 68 to 105 cN/dtex, respectively (Figure 4d). The tensile strength values of these fibers were higher than other pure chitin fibers prepared from other solvents, such as in xanthate (0.71-1.20 cN/dtex),54 formic/dichloroacetic acid (0.60-1.40 cN/dtex),55 and N, N'-dimethylacetamide/lithium chloride (0.62-1.90 cN/dtex).56 14


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Moreover, the tensile strength of our chitin fibers was also higher than most of the natural polysaccharides-based other fibers, such as agar/agarose fibers (0.44-0.69 cN/dtex),57 alginate fibers (1.16-1.93 cN/dtex),58 carrageenan fibers (0.73-0.75 cN/dtex)59 etc, but lower than cellulose fibers (2.0-3.5 cN/dtex) fabricating by the same method.33 It was worth noting that the tensile strength of chitin fibers fabricated from the same NaOH/urea system but absence of nanofibers was in the range from 0.75 to 1.36 cN/dtex with an increase of the drawing ratio.42 Importantly, the tensile strength of the chitin fibers in the wet state was 1.0-1.5 cN/dtex, similar with that of modal, a new type of cellulose fiber (1.5 cN/dtex in the wet state),60 demonstrating the fibers were good candidate as biomedical materials, especially for tissue engineering. This made the commercialization possibility of the chitin fibers (Figure 4c). It was anticipated that the tensile strength of the chitin fibers could be further enhanced by optimization in the industrialized production facility. In our findings, the strong self-aggregation force10 between chitin chains and bundles driven by numerous hydrogen bonds sustained the nanofibers and densely aligned structure, leading to the enhancement of their tensile strength. Indeed, the enhancement of the mechanical properties endowed the fibers with the possibility of wider applications in various fields. Furthermore, the fabrication of strong fibers via physical dissolution and physical regeneration in aqueous medium has been demonstrated to be environmentally-benign (Table 1), suggesting a “green” and sustainable avenue for the potential commercialization of the chitin fibers.

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Table 1. Mechanical properties of regenerated chitin fibers. Solvents used

Coagulating bath

σ (cN/dtex)

ε (%)

Ref.

NaOH/urea

phytic acid/Na2SO4

1.66-2.33

5.2-8.6

This work

NaOH/urea

H2SO4

0.75-1.36

12-24

42

chitin xanthate

H2SO4/Na2SO4/ZnSO4

0.71-1.20

--

54

formic acid /dichloroacetic acid

ethyl acetate/water

0.60-1.40

2.7-4.3

55

DMAc-LiCl

methanol or DMAc/H2O

0.62-1.90

--

56

concentrated H2SO4

alkali or alcohols

2.20

--

61

trichloroacetic acid /dichloromethane

--

2.30

--

62

Figure 5. The morphology and structure of NRVM cultured with chitin fiber. (a) Bright field images and (b) confocal image showed NRVM cultured with chitin fibers in hydrogel for 7 days, dotted lines represent the contours of chitin fibers. (c) and (c’) show sarcomeric α-actinin (SAA) in red, nuclei (DAPI) in blue. Scale bar: (a, c) 20 μm, (b) 50μm, (c’) 10μm.

Chitin Fiber Support Growth of Ventricular Myocytes An idea scaffold is required to be nontoxic, biocompatible and safety. To evaluate the potential application of the chitin fibers in biomedical and regeneration medicine, the customized chitin fiber 16


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scaffold was tested to culture the neonatal rat ventricular myocytes (NRVM) (Figure S6). Figure 5ac’ shows the bright field images and immunofluorescence staining images of NRVM cultured with chitin fiber scaffold in fibrinogen/matrigel hydrogel. Clearly, the chitin fiber scaffold exhibited enough mechanical strength to support the cardiac tissue (Figure 5a, b). Interestingly, NRVM on this scaffold exhibited spontaneous beating (Video S1) and clear sarcomere structure after being cultured for 7d. This indicated that the chitin fiber scaffold was not only robust, and also nontoxic and biocompatible for the cardiac tissue. Furthermore, nonwoven prepared by chitin fibers could shorten the wound healing time from 16 days of commercial gauzes to 12.5 days, as a result of the inherent antibacterial properties of chitin and its ability to regulate inflammatory mediators.42 These results demonstrated the chitin fibers could be a potential supporting material for ventricular myocytes and displayed potentials in the tissue engineering. It is not hard to imagine that the chitin fiber retaining the intrinsic structure and character of native α-chitin is a good candidate for the biomedical materials.

17



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Figure 6. Degradation test. (a) Digital pictures (top) and SEM images (bottom) of the chitin fibers after degrading for 2, 5, 8 and 11 days in soil; (b) Digital pictures (top) and SEM images (bottom) of the chitin fibers after degrading for 5, 11, 17 and 30 days in vitro; (c) Dependences of weight loss of the chitin fiber on degradation time in soil and in vitro.

Biodegradation Behaviors To protect environment, biodegradability may be a great advantage of the natural polysaccharide-based materials.63 Thus, the chitin fibers were buried in soil to perform the biodegradation test (Figure 6). The chitin fibers remained its shape after 2 days (Figure 6a), whereas they were biodegraded by the microorganisms in the soil, leading to the loose fractionlet after 5 days, corresponding with the rougher surface as shown in the SEM image. Only a few particle-like sample appeared after burying in soil for 8 days, indicating a good biodegradability of the chitin fibers. This phenomenon can be explained that the chitin was attacked and digested by the microorganisms and thus was gradually biodegraded in the soil.64 From the data in Figure 6c, the complete biodegradation time of the chitin fiber could be extrapolated to be approximately 22 days in soil at average environment temperature of 30 oC, which was faster than the chitin sponge (32 days) reported in our laboratory,64 possibly due to the relatively high temperature and larger contact area of the chitin fibers with soil than the sponge. Thus, the chitin fibers could act as substitutes of plastics in many fields, such as daily necessities, textile, diapers, carpets and ropes,65 in which the complete biodegradation is very important. In that case, the chitin fiber based products discarded into the soil and sea after using will be completely degraded, which would not cause any pollution. Therefore, the chitin fibers constructed from the renewable resource via “green” process had excellent biodegradability, showing promising applications in wide fields for the virtuous circle of the world. 18


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It is well known that bioabsorbable medical devices refer to those made of biodegradable and absorbable materials in the biological environment, which can be biodegraded and absorbed in the biological environment in vivo. Their application fields mainly focus on absorbable surgical sutures, hemostasis medical devices, internal fixation of fracture, and absorbable adhesion proof product, etc., so they do not need to carry out secondary surgery in clinical use. Thus, the in vitro degradation of the chitin fiber in phosphate-buffered solution in the presence of lysozyme at 37 oC were investigated. As shown in Figure 6b, the chitin fibers were destroyed at 11 days, displayed fractionlet sample at 17 days and remained a few particle-like sample after immerging in PBS for 30 days. The degradation rate of chitin fibers raised with an increased concentration of lysozyme (Figure S7). From the results of biodegradation test (Figure 6c), the complete biodegradation time of the chitin fiber in vitro could be extrapolated to be approximately 34 days, much shorter than polymer fibers, and similar with other reported chitin fibers (about 30 days).66 In our laboratory, it has been already confirmed that chitin bioplastic exhibited good in vivo biodegradability.13 Therefore, the chitin fibers possessed the attractive advantages including excellent biocompatibility as well as biodegradability both in nature and in living organisms. As shown in Table S3, the good mechanical properties and biodegradability of the chitin fibers made it potential in biological application, while more concrete medical applications of the chitin fibers should be developed.67-70 In view of these advantages, the chitin fibers constructed from the seafood wastes via “green” process not only show promising applications as substitutes to the non-degradable daily necessities, but also illustrate potential utilizations in the fields of bioabsorbable medical devices. This work opened out a new avenue for more wide applications of the chitin as sustainable polymer material.

CONCLUSIONS 19



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The chitin fibers with nanofibril structure were fabricated successfully from the chitin solution via a “bottom-up” approach on a lab-scale wet-spinning machine. In our findings, the chitin molecular chains dissolved in NaOH/urea aqueous solution could self-aggregate easily in a parallel manner to form nanofibers in the phytic acid/Na2SO4 coagulant bath. The dried chitin fibers were composed of nanofibers, and displayed excellent tensile strength. With an increase of the draw ratio from 1.0 to 1.8, the crystal index and degree of orientation increased slightly, whereas the tensile strength and Young’s modulus of the chitin fibers enhanced significantly. The strong self-aggregation force between chitin chains and bundles driven by numerous hydrogen bonds sustained the nanofibers and densely aligned structure. Furthermore, the chitin fibers retained the intrinsic characteristics of native α-chitin, and displayed good biocompatibility towards NRVA cells by promoting their adhesion and growth. Biodegradation tests demonstrated that the chitin fibers can be completely biodegraded in soil and in vitro. This work provided a new avenue to construct strong chitin fibers with biocompatibility and biodegradability, to the benefit of both avoiding secondary surgery in clinical use and protecting environment.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: Schematic diagram and photographs of the wet spinning machine; SEM images of the cross-sectional structures of different chitin fibers; SEM images of the surface structures of the chitin fibers in the dry state; Chemical structure of chitin; Stress-strain (σ-ε) curves of chitin fibers; The process of culturing NRVM with chitin fiber in Matrigel/fibrinogen hydrogel; Degradation test in vitro under different concentration of lysozyme; Spinning parameters of different chitin fibers; Crystallinity, 20


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degree of orientation, degree of acetylation and mechanical properties of the chitin fibers; Mechanical properties, advantages and disadvantages of different fibers for biological applications (PDF) Video of the spontaneous beating of cells cultured on the chitin scaffold (AVI)

AUTHOR INFORMATION Corresponding Authors *E-mail: [email protected] (L. Zhang) *E-mail: [email protected] (Q. Fu) *E-mail: [email protected] (A. Lu) Author Contributions # K. Zhu and H. Tu contributed equally to this work. ORCID Lina Zhang: 0000-0003-3890-8690 Ang Lu: 0000-0001-6457-8264 Notes The authors declare no competing financial interest.

ACKNOWLEDGEMENTS This work was supported by the Major Program of National Natural Science Foundation of China (21334005, 51421061, 51573143), the Major International (Regional) Joint Research Project (21620102004),

International Cooperation and Exchange of the

National

Natural

Science

Foundation of China (21811530006), the National Natural Science Foundation of China (20874079, 31871496), and the Fundamental Research Funds for the Central Universities (2042018kf0042). 21



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Mechanically Strong Chitin Fibers with Nanofibril Structure

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