Robust Anisotropic Cellulose Hydrogels Fabricated via Strong Self

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Robust Anisotropic Cellulose Hydrogels Fabricated via Strong Selfaggregation Forces for Cardiomyocytes Unidirectional Growth Dongdong Ye, Pengcheng Yang, Xiaojuan Lei, Donghui Zhang, Liangbin Li, Chunyu Chang, Pingchuan Sun, and Lina Zhang Chem. Mater., Just Accepted Manuscript • DOI: 10.1021/acs.chemmater.8b01799 • Publication Date (Web): 20 Jul 2018 Downloaded from http://pubs.acs.org on July 21, 2018

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Chemistry of Materials

Robust Anisotropic Cellulose Hydrogels Fabricated via Strong Self-aggregation Forces for Cardiomyocytes Unidirectional Growth Dongdong Ye,† Pengcheng Yang,‡ Xiaojuan Lei,† Donghui Zhang,‡ Liangbin Li,§ Chunyu Chang,†,* Pingchuan Sun,ǁ and Lina Zhang†,* †

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

‡

College of life science, Hubei University, Wuhan 430062, China

§

National Synchrotron Radiation Lab and College of Nuclear Science and Technology,

University of Science and Technology of China, Hefei 230026, China ǁ

Key Laboratory of Functional Polymer Materials of Ministry of Education and College of

Chemistry, Nankai University, Tianjin 300071, China ABSTRACT The development of a facile and fast method to construct anisotropic hydrogels with the ability to induce unidirectional growth of cells remains challenging. In this work, we demonstrated anisotropic cellulose hydrogels (ACHs) that are composed of nanoscale aligned nanofibers by dissolving cotton liner in alkali/urea aqueous solution. Based on the directionally controlling the architecture of cellulose chains with a facial pre-stretching strategy in chemical gel state and locking the highly ordered nanostructure through the formation of close physical networks via strong self-aggregation forces among neighboring cellulose nanofibers, ACHs, combing with a long-range aligned structure, entirely differential mechanical performances along the parallel and perpendicular directions of the hydrogel orientation and optical birefringence, were constructed. The aggregation of hydrogen bonds in anisotropic and isotropic hydrogels are of significant difference, confirmed by nuclear magnetic resonance technology. Importantly, ACHs with microgroove-like structure, promote the adhesion and orientation of cardiomyocytes. Our work 1 ACS Paragon Plus Environment

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demonstrated the bottom up fabrication of polysaccharide-based hydrogels with anisotropic structure and properties, paving the way to potential apply them in cardiomyocytes in vitro culture system. INTRODUCTION Hydrogels possess water-rich structures similar to biological tissues, and therefore, they have been regarded as promising scaffolds in biomedical fields.1,2 Biological systems adopt anisotropic structures with hierarchically integrated building units, and thus, hydrogels with highly ordered structures, unique properties and wide applications in sensors, actuation and anisotropic cell culture have gained widespread interest in material science fields.3-5 To obtain a long-range aligned structure in hydrogels, various strategies, such as electrical field alignment,6,7 magnetic field alignment,8-10 strain alignment,11-14 directional freezing,15,16 ion alignment,17,18 selfassembly19-21 and drying in confined conditions22 have been employed. External fields are generally used to induce the temporary orientation of nanofillers or molecular chains prior to the subsequently complex polymerization or gelation process. For example, ionic crosslinking has been used to lock aligned macromolecules through Ca2+/alginate23 or Fe3+/-COO- interactions13 to obtain permanently oriented hydrogels, but these methods are time-consuming. Thus, the development of a facile and fast method to construct anisotropic hydrogels with the ability to induce unidirectional growth of cells remains challenging.24 Considering the fact that cardiovascular disease is the most significant cause of morbidity and mortality in the world and due to the shortage of functional cardiomyocytes for drug discovery and regenerative therapy, a highly organized homologation arrangement is critical for maintaining cardiomyocyte function, such as their electrophysiological properties and contractility. Although various substrates have been developed for cardiomyocyte in vitro culture,25-27 biocompatible scaffolds combining highly oriented structure with excellent 2 ACS Paragon Plus Environment

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mechanical properties are still seldom reported. Cellulose is the most abundant naturally occurring macromolecule, and has received renewed interest due to its wide application via “green” conversion.28 Hu et al reported a high-performance densified wood consisting of wellaligned cellulose nanofibers, resulting from the formation of packed hydrogen bonds among neighboring nanofibers.29 Gong et al revealed that supramolecular interactions such as hydrogen bonds through the summation of a directionally controlled hierarchical architecture can be very strong and can stabilize the oriented structure.22,30 The stiff molecular chains and close packing through the numerous hydrogen bonds of cellulose, namely, the existence of strong selfaggregation forces in bulk cellulose, make its dissolution a very difficult process. Benefitting from the cellulose solvent-alkali/urea aqueous system of our research group,31 herein, we designed and fabricated anisotropic cellulose hydrogels with diversified shapes through a novel and facile strategy, combining facile pre-stretching and acid treatment processes. Particularly, our anisotropic polysaccharide-based hydrogels with a microgroove-like oriented structure, as a first example, was used for the adhesion, growth and maturation of cardiomyocytes, revealing their promising application in using as a new in vitro cell maintains. EXPERIMENTAL SECTION Materials. Cellulose (cotton linter pulp) with α-cellulose content beyond 95% was provided by Hubei Chemical Fiber Group Ltd. (Xiangfan, China). The cotton linter pulp was used after complete drying under a vacuum at 60 °C and without further purification. The viscosity-average molecular weight of the cotton linter pulp in cadoxen was determined as 9.6×104 (degree of polymerization, DP=600) according to the Mark-Houwink equation [η] (mL g-1) = 3.85×10-2 (Mw)0.76 and using an Ubbelohde viscometer at 25 °C. LiOH·H2O, urea, concentrated sulfuric acid and epichlorohydrin (EPI) were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). All of the reagents were used as received unless otherwise noted. 3 ACS Paragon Plus Environment


Loosely chemically cross-linked cellulose gels (LCGs) preparation.

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LCGs were fabricated

according to our previously reported work32. Briefly, a cellulose solution (6 wt%) was prepared by dissolving cotton linter pulp in a pre-cooled 4.5 wt% LiOH/15 wt% urea aqueous solution. Certain amounts of EPI corresponding to EPI:AGU (anhydroglucose unit) =1:4.64, 1:3.09, 1:2.32, 1:1.86 and 1:1.55 were added to the cellulose solution while stirring at 0 °C for 2 h. Then, the viscous cellulose solution was poured into molds after removing air bubbles via centrifugation and kept at 5 °C for 8 h. After removing from the molds, LCGs were obtained and coded as LCG-1 to LCG-5. Preparation of anisotropic cellulose hydrogels (ACH). LCGs were pre-stretched to certain DRs and immersed in 5 wt% aqueous sulfuric acid within one minute for the formation of physically cross-linked networks with nanofibers and to fix the highly oriented structure in the hydrogels through hydrogen bonds. ACHs with permanent shapes were obtained after thoroughly washing the hydrogels with deionized water.

Scheme 1. Preparation of anisotropic cellulose hydrogels (ACHs). First, the loosely chemical cross-linked gel (LCG), fabricated from cellulose solution, was pre-stretched by external force, and the hydrogel network aligned in the stretching direction to form a temporarily oriented 4 ACS Paragon Plus Environment

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structure. Then, a relatively close-packed architecture including hybrid crosslinking networks formed in the acid solution driven by the strong self-aggregating force of the cellulose itself, leading to permanent locking of the orientation structure of the hydrogels. Neonatal rat ventricular myocyte (NRVM) isolation and culture. Cardiomyocytes from neonatal (1 to 2-day-old) Sprague-Dawley rats were isolated using a Neomyts kit (Cellutron, NC6031). The resulting cell suspension was pre-plated in 0.1% gelatin-coated 10-cm Petri dishes for 2-h intervals to enrich the cardiomyocytes from the adherent cells (non-myocytes). The wet sterilized hydrogels were cut into 0.5 mm×0.5 mm×0.2 mm (length, width, and height) sheets and coated with fibronectin (Sigma, F0895) in 24-well plates. Then, 500 µL of a fresh NRVM suspension at 1×106 cells/mL was seeded on the hydrogels at 37℃ for 5 d. The NRVMs were cultured in medium consisting of Dulbecco’s modified Eagle’s medium (DMEM) with 4.5 g/L glucose, 5% certified fetal bovine serum (FBS), 1% streptomycin/penicillin and 50 µg/µL ascorbic acid; 20 µM cytarabine (Sigma, c1768) was added after the first 24 h to inhibit proliferation of non-myocytes and to preserve the initial purity of the NRVMs. The cells grown on the hydrogels were stained by MitoTracker™ Red CMXRos (Life Technologies, M7512), Sarcomeric, α-actinin (Sigma, A7811') and DAPI. Confocal images were taken using a Zeiss LSM710. Measurement. AFM images of the hydrogels were recorded using a CypherTM S (Asylum Research) microscope in a fluid environment by using silicon nitride probes (SNL-10, BRUKER) with a tip radius of 2 nm, a spring constant of 0.35 N/m and a resonance frequency of 60 kHz. SEM images of freeze-dried ACHs were taken with an FE-SEM (Zeiss, SIGMA, Germany) operating at an acceleration voltage of 5 kV. Tensile testing of the hydrogels was performed on a universal tensile-compressive instrument (Model 5576, INSTRON Instrument, USA). Hydrogel specimens measuring 50-mm long and 10-mm wide were stretched at a speed of 5 mm/min. Cyclic loading and unloading tests, using the same specimens at an equal test speed, were 5 ACS Paragon Plus Environment


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performed in subsequent trials immediately after the initial loading. 2D SAXS measurements of the hydrogels were obtained using an in-house micro X-ray source (Incoatec, GmbH) with Cu Kα radiation (λ = 0.154 nm). The scattering signal was collected by a multiwire proportional chamber detector (Bruker, Hi-STAR) with a resolution of 487 × 619 pixels (pixel size of 105 µm). The sample-to-detector distance was fixed at 3115 mm. 2D WAXS was carried out on an Xray scattering station with a Mar 345 image plate (3072 × 3072 pixels with a pixel size of 150 µm) as the detector and with a wavelength of 0.154 nm. The sample-to-detector distance was calibrated to be 253 mm. The 2D scattering images were analyzed with Fit2D software from the European Synchronization Radiation Facility. Herman’s orientation parameter (fc) was calculated from the azimuthal-integrated intensity distribution curves of the SAXS and WAXS patterns with the following equations:

fc =

3 < cos 2φ > − 1 2

∫ < cos φ >= 2

(1)

π /2 0

I (φ )cos 2φ sinφ dφ

∫

π /2 0

I(φ )sinφ dφ

(2)

where is the azimuthal angle, and ( ) is the 1D intensity distribution along with . The value of is calculated by integrating the intensity of the specific 2θ diffraction peak along using the aforementioned equation. The X-ray diffraction (XRD) patterns of the aforementioned cellulose hydrogels were recorded on a Riguka Smartlab 9k diffractometer operated at 45 kV, 200 mA in reflection mode for Cu Kα (λ=0.154 nm) with a scan speed of 2°/min and a step size of 0.05° in 2θ. Solid-state nuclear magnetic resonance (NMR) experiments were performed on a Varian Infinity Plus wide-bore (89 mm) NMR spectrometer operating at a 1

H frequency of 399.7 MHz and a

13

C frequency of 100.5 MHz. A conventional 5-mm double-

resonance HX CP/MAS NMR probe was used, and D2O-exchanged cellulose hydrogels were 6 ACS Paragon Plus Environment

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placed in a 5-mm zirconia PENCIL rotor with a well-sealed cap and spacer. The magic angle spinning (MAS) was automatically controlled at 6 kHz within ± 2 Hz with a MAS speed controller. CPMAS, DPMAS and cross-polarization with the nuclear Overhauser enhancement effect (CPNOE) experiments, which are used to obtain the rigid, mobile and total (rigid and mobile) components, were carried out at room temperature. Heteronuclear decoupling during the acquisition period was achieved by SPINAL-64 irradiation with a radio frequency strength of ~80 kHz. The 13C chemical shift was referenced to external hexamethylbenzene (HMB, 17.3 ppm of CH3). The background signal of the rotor at 111.3 ppm in the 13C DPMAS and CPNOE spectra should be ignored in Figure 2c. RESULTS AND DISCUSSION Construction of anisotropic cellulose hydrogels (ACHs) via self-aggregation force

Figure 1. Photograps of LCG (a), pre-stretched LCG (b) and ACH under polarized light (c). DR: draw ratio, and DR=L/L0; A: analyzer; P: polarizer.

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Anisotropic cellulose hydrogels composed of nanoscale aligned nanofibers were fabricated from cellulose in alkali/urea aqueous solution via a bottom-up method. Here, we utilized the strong self-aggregation trend of cellulose itself to serve as the driving force for the construction of highly oriented ACHs. Scheme 1 displays the formation of ACH with a highly ordered structure. First, loosely cross-linked gel (LCG) was synthesized by reacting cellulose with a small amount of epichlorohydrin (EPI) in a LiOH/urea aqueous solution. The LCG containing solvent (LiOH-urea) could be easily pre-stretched under external force to form an oriented structure, and its original shape could be immediately recovered after removal of the force (Figure 1a, b; Movie S1). Subsequently, the pre-stretched LCG with temporarily aligned networks was immersed in sulfuric acid aqueous solution to rapidly fix the oriented structure and shape of the hydrogel within one minute. Acid treatment could thoroughly destroy the alkali/urea solvent shell on the cellulose chains to easily remove the solvent molecules from the network, and thus, the realigned orientation structure and the designed shape of the hydrogels could be permanently locked through intermolecular hydrogen bonds between cellulose. After washing with deionized water, permanently oriented cellulose hydrogels with pre-stretching draw ratios (DR=X) and optical birefringence were obtained and coded as ACH-X (Figure 1c).


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Figure 2. Differences in structure and properties between isotropic and anisotropic cellulose hydrogels. (a) SEM images of the cross-sections of the isotropic and anisotropic hydrogels. (b) Photographs of triangle-shaped isotropic (top) and anisotropic (bottom) hydrogels under visible light and polarized light. Note: The background of photographs is the school badge of Wuhan University, used with permission. (c) Results of

13

C CPMAS,

13

C DPMAS and

13

C CPNOE

NMR of the isotropic and anisotropic cellulose hydrogels. Note: * represents the background signal of the rotor. Moreover, filed emission scanning electron microscopy (FE-SEM) images of the hydrogel cross-section (Figure 2a) indicated that the anisotropic hydrogel consisted of oriented submicronscale channels composed of nanofibers, whereas the isotropic hydrogel exhibited a porous structure, demonstrating that the aggregation structures (chain packing, close alignment and orientation) of LCG samples changed significantly from isotropic to anisotropic after prestretching and acid treatment. The morphology of ACH was similar to that of wood, where thousands of nanofibrils oriented in the cell walls along the tree growth direction, as shown in the cross-section and surface SEM images (Figure S1). This demonstrated that the self-aggregation force sustained the rearranged orientation structure of ACHs. What’s more, the photographs of 9 ACS Paragon Plus Environment


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isotropic and anisotropic hydrogels under both visible and polarized light (Figure 2b) show their isotropic and anisotropic optical properties, respectively. These hydrogels were transparent under visible light. Birefringence was only exhibited in the anisotropic hydrogel, while no birefringence appeared in the isotropic cellulose hydrogel, suggesting that the highly aligned structure in anisotropic hydrogel dominated the optical anisotropy. To further investigate the structure of the anisotropic hydrogels and to detect the formation of self-aggregation forces within the cellulose, nuclear magnetic resonance (NMR) technology was employed. Figure 2c shows the polarization magic angle spinning (CPMAS), (DPMAS) and

13

13

C cross-

13

C direct-polarization magic angle spinning

C cross-polarization with nuclear overhauser enhancement effect (CPNOE)

spectra of anisotropic and isotropic hydrogels, respectively. In the

13

C CPMAS spectrum of

anisotropic hydrogel, the appearance of the shoulder at 103 ppm for C1, the increase in resolution at 88 ppm for C4, and the split of the peak at 62 ppm for C6 indicated that the hydrogen bonds became arranged and the crystallinity of the hydrogel increased, compared with those of isotropic hydrogel.33 For the 13C DPMAS spectra, obvious changes in characteristic peaks (blue region) for rigid components of the anisotropic hydrogels were observed, but the mobile components that interacted with water changed slightly. Furthermore, the results of the

13

C CPNOE spectra also

confirmed that nanofibrils (rigid components) formed in the anisotropic hydrogel through the self-aggregation force of cellulose chains.34 Therefore, the NMR results strongly supported that the aggregation of hydrogen bonds in anisotropic and isotropic hydrogels are obviously different. In view of the above results, the formation mechanism of ACHs could be described as follows: i) the removable cellulose rigid chains and bundles in the chemical gel temporarily rearranged along the pre-stretching direction; ii) the highly ordered structure of the pre-stretched gel was locked in acid solution through the strong self-aggregation force of cellulose (mainly the


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hydrogen bonding interactions); and iii) the self-aggregation force played an important role in the formation of tough anisotropic cellulose hydrogels with highly ordered structures. Highly oriented nanostructure of anisotropic cellulose hydrogels The morphology, structure and degree of alignment of ACHs were characterized by atomic force microscopy (AFM), FE-SEM, and X-ray scattering techniques in a synchronization radiation facility. As shown in Figure 3a, ACH-1.0 without pre-stretching treatment displayed a homogeneous morphology and porous structure, whereas anisotropic cellulose hydrogels gradually tended to form oriented structures along the pre-stretching direction with increased DR values. For example, ACH-1.4 exhibited a porous network structure, which slightly oriented along the pre-stretching direction (Figure S2). Once DR increased to 1.8, the cellulose nanofibers were further rearranged and reoriented along the pre-stretching direction in ACH-1.8 (Figure S3). Particularly, the cellulose nanofibers (mean diameter: 39 nm) could align closely, leading to a tendency to assemble and stack with each other into more compact submicrofibrils (mean diameter: 578 nm) as the DR increased to 2.2 (Figure S4). More importantly, highly aligned nanofibers and denser structures appeared in ACH-2.6, further confirming that acid induced the conversion of the aggregation structure in the pre-stretched gel.



Figure 3. Morphology and orientation analysis of the cellulose hydrogels with various DRs. (a) Representative AFM height images of cellulose hydrogels under pre-stretching DRs of 1.0, 1.4, 2.2 and 2.6, respectively. (b, c) FE-SEM images of surfaces and cross-section of ACH-2.6 at different magnifications. (d) 2D SAXS patterns, and e) 2D WAXS patterns of ACH-1.0 to ACH2.6 hydrogels. To clarify the morphology of ACH-2.6, FE-SEM images of the surfaces of freeze-dried samples are shown in Figure 3b. Oriented cellulose nanofibers could be observed in the surface of the hydrogel, consistent with the AFM results. Moreover, densely well-aligned cellulose 12 ACS Paragon Plus Environment

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nanofibers also appeared in the cross-section of freeze-dried ACH-2.6 (Figure 3c). The cellulose hydrogels exhibited aligned hierarchical fibrous structures with submicro-scale hole-like aisles, which were occupied with water. The pre-stretching DR values of LCGs not only effected the morphology of ACHs but also played an important role in their water content, deformation degree after absorbing water, and crystallinity. The water content of ACHs gradually decreased from 91.01% ± 0.31% to 85.01% ± 0.54% with an increase in DR from 1.0 to 2.6 (Figure S5). Besides, the deformation tests of isotropic hydrogel (ACH-1.0) and anisotropic hydrogels (including ACH-1.4, 1.8, 2.2 and 2.6) in the parallel and perpendicular of hydrogel orientation directions after absorbing water were conducted (Figure S6), and it demonstrated that ACH-1.0 exhibited a nearly homogeneous swelling behavior along parallel and perpendicular direction, while the anisotropic hydrogels exhibited a several fold difference in the transverse and longitudinal swelling strains. Especially for ACH-2.6 with denser highly ordered structure, the difference in two directions reached 9.61-folds on the one hand, but on the other hand anisotropic hydrogels exhibited smaller deformation degree (1.0) displayed elongated longitudinal 2D patterns, 13 ACS Paragon Plus Environment


suggesting the presence of aligned nanofibers in the cellulose hydrogel. In particular, the sharpest elongated SAXS pattern of ACH-2.6 was observed, indicating that highly aligned cellulose nanofibers were tightly stacked in the hydrogel, resulting in its anisotropic morphology. Moreover, wide angle X-ray scattering (WAXS) technology was also employed to characterize the ordered structure of ACHs (Figure 3e). For ACH-1.0, a nearly uniform diffraction pattern appeared at all azimuthal angles on the (110) scattering plane, suggesting an isotropic structure. With an increase in DR from 1.0 to 2.6, the equatorial arcs of patterns tended to become clearer, revealing the anisotropic characteristics of the hydrogels with the aligned nanofibers. We should emphasize that the WAXS patterns were obtained by removing the backgrounds of water and air because the pattern of cellulose was overlapped with the patterns of water and air (Figure S8). As the DR increased from 1.0 to 2.6, the Herman’s orientation parameter (fc) of the cellulose hydrogels gradually rose from 0.031 to 0.84 according to SAXS and from 0.026 to 0.82 according to WAXS (Figure S9). The high fc values of the ACHs further demonstrated the existence of a highly aligned architecture along the pre-stretching direction, strongly confirming that the self-aggregation force of the cellulose drove the formation of highly ordered anisotropic cellulose hydrogels. This is very different from flexible polymers, which could not form intraand intermolecular hydrogen bonds to maintain the rigid chains and oriented structure of materials. Mechanical properties of anisotropic cellulose hydrogels To clarify the relationship between the designed structure and properties of hydrogels, the mechanical properties of LCGs and ACHs were evaluated. The fracture strains of LCGs could be controlled by changing the molar ratio of EPI/AGU in the crosslinking reaction (Figure 4a). The tensile stress-strain curves of LCGs exhibited high stretchability under a small tensile force of kilopascals (Figure 4a; Movie S1), and LCG-4 with epichlorohydrin (EPI)/ anhydroglucose unit 14 ACS Paragon Plus Environment

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(AGU) molar ratio of 1:1.86 displayed the highest fracture strain (~180%) and extremely low dissipated energy (10.04 kJ m-3). Importantly, the inapparent hysteresis of LCG-4 was observed by the cyclic stress-strain curves under fixed deformation from 40% to 180% with a constant step to maximum stretching (Figure 4b). Therefore, LCG-4 was selected for the preparation of the following ACH hydrogels. Figure 4c, d shows the effect of the DR value on the mechanical properties of the ACHs along parallel (//) and perpendicular (⊥) directions. The tensile strength and fracture energy of the isotropic hydrogel (ACH-1.0) were 0.86 MPa and 0.49 MJ m-3, respectively, which were far higher than those of LCGs (Figure 4a, c and e), suggesting that the introduction of physical crosslinking in the networks significantly improved the mechanical properties of the hydrogel. Furthermore, the mechanical strength of ACHs enhanced with an increase in DR values along the parallel direction (Figure 4c). More importantly, the tensile strength, elastic modulus, and fracture energy of ACH-2.6 in the parallel direction dramatically increased, reaching 7.98 MPa, 37.93 MPa, and 1.08 MJ m-3, respectively; however, the fracture strain decreased rapidly to 26.4% (Figure 4f).



Figure 4. Mechanical properties of the cellulose hydrogels. (a) Tensile stress-strain curves of LCGs with differential dosage of chemical crosslinker, showing that LCGs could be easily stretched to form oriented structures under a small tensile strength. (b) Representative tensile stress-strain curves of LCG-4 during loading-unloading cycles of varying strains, demonstrating the high elasticity of LCG-4 and its rapid recoverability after removal of the external force. (c, d) Tensile stress-strain curves of ACHs with different DRs along the parallel and perpendicular directions, indicating significant differences in stiffness and flexibility. (e, f) Elastic modulus and fracture energy of the ACHs along the parallel and perpendicular directions. It was noted that in the perpendicular direction, the tensile strength and elastic modulus of ACHs were much lower than those in the parallel direction, whereas their fracture strains and fracture energies enhanced from 144.1% to 316% and from 0.49 to 0.83 MJ m-3, respectively, as the DR value increased from 1.0 to 2.6 (Figure 4d, f). To visually differentiate their stiffness and stretchiness, photographs of the anisotropic hydrogels under polarized light during the tensile process along the // and⊥ directions were recorded (Figure S10; Movie S2). ACH-2.6-// could retain its shape to overcome gravity, and displayed extremely high stiffness. However, ACH-2.6⊥was too soft and flexible to retain its shape under gravity. The anisotropic hydrogels exhibited bright birefringence under polarized light and a significant distinction of fracture strain in the parallel and perpendicular directions. The great differences in the stiffness and stretchiness of the ACHs in the two directions indicated their anisotropic mechanical behaviors. The mechanical anisotropy of ACHs along the parallel and perpendicular directions could be explained as follows: i) Their ultrahigh tensile strength in the parallel direction was attributed to the strong intermolecular hydrogen bond interactions between cellulose chains and nanofibers in the highly compact form, which were favorable for the removal of defects in the hydrogel; ii) In the perpendicular direction, the densely packed cellulose network could be easily stretched to a loose


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state by breaking the hydrogen bonds, thus forming a new alignment structure and leading to the high fracture strain of the hydrogel. Design of anisotropic cellulose hydrogels with diversified shapes and structure Importantly, ACHs with diversified shapes were successfully fabricated by acid treatment of pre-stretched LCGs after twisting or releasing (Figure 5). For example, ACHs with microgroovelike structures along the perpendicular direction could be generated via releasing the prestretched LCG in diluted acid solution (Figure 5a). When a pre-stretched chemical gel with temporarily aligned networks was immersed in sulfuric acid aqueous solution and simultaneously removed external force. The external structure of chemical gel firstly contacted with acid and became a relatively hard thin layer through the formation of dual crosslinking networks, whereas the internal domain of gel was still soft and tended to shrink after removal of force, leading to the formation of a wrinkled structure.35 As shown in Figure 5b-d, ACHs with a wrinkled structure had almost parallel-distributed microgrooves along the perpendicular direction on the surface of the cellulose hydrogel, and the ridge and valley regions exhibited distinguishing polarization phenomena under polarized light, confirming the microstructural differentiation among these regions. The morphologies of the ridge and valley regions of the ACHs were characterized by FE-SEM, and their regions exhibited significantly structural differences, as mentioned above (Figure 5e). The long-scale aligned pattern was still observed in both ridge and valley regions, and the morphology in the ridge region became more dense and rough, while the valley region showed a loose and smooth structure. Based on the highly aligned structure and the distinctive morphology in ACHs, these cellulose hydrogels might have great potential applications for improving of cell adhesion in tissue engineering.



Figure 5. (a) Photographs of diversified ACHs constructed with pre-stretched LCGs and acid treatment. (b, c) Photographs of ACHs with microgroove-like structures constructed via prestretching and releasing of the LCGs in diluted acid solution under visible light (top) and polarized light (bottom). (d) Representative morphology of ridge and valley regions in the ACH surface. A: analyzer, P: polarizer. (e) FE-SEM images of ACHs with microgroove structures. Cellulose hydrogels induced unidirectional growth of cardiomyocytes Considering the fact that anisotropic cellulose hydrogels with low roughness (20 nm) cannot adhere neonatal rat ventricular myocyte (NRVM) (Figure S11), the microgroove-like cellulose hydrogel with average roughness of 132 nm on ridge regions (Figure S12) was used to support NRVM growth and maturation in vitro so as to verify the potential biomedical applications of ACHs, as shown in Figure 6. While cultured on the hydrogel, the NRVMs attached on both the 18 ACS Paragon Plus Environment

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ridge and valley-like region of hydrogel, and aligned along the direction of the hydrogel orientation (Figure 6a-d). The cultured NRVMs exhibited a mitochondrial enrichment (Figure 6e and Figure S13), the clear sarcomere structure (Figure 6d and Figure S14) and spontaneous beating (Movie S3). The average sarcomere length was 1.79 ± 0.14 µm (Figure S15), which is comparable with the cultured NRVM (1.94 ± 0.07 µm),36, 37 and similar with the isolated adult rat myocytes (1.71 ± 0.10 µm).38 These results confirmed the physiological activity and structural maturation of the cultured cardiomyocytes. It was worth noting that the orientation of the cellulose nanofibers in the hydrogel was in fact perpendicular to the orientation of the microgrooves. Usually, the cells filopodium can extend and adhere on the nanofibers of the biomaterials.39 Due to their microgroove-like structure, ACHs could promote well the adhesion and orientation of cardiomyocytes, resulting in the ability of the cardiomyocytes to connect with each other and contract together (Figure 6f, Movie S3). Compared with irregularly oriented cardiomyocytes traditionally cultured on coverslips (Figure 6g), the anisotropic cellulose hydrogel with microgroove-like structure significantly induced cardiomyocyte morphological orientation (Figure 6h), according to the statistics of their relative directions (Figure 6i). These results confirmed that the unique orientation structure of the hydrogel could promote the adhesion and arranged orientation of cardiomyocytes, suggesting potential functions for improving the electron propagation and contractility of cardiomyocytes maintained in vitro. This work provides new applications of microgroove-like anisotropic hydrogels in cardiomyocytes in vitro maintains.



Figure 6. Adhesion and oriented growth of NRVMs on the anisotropic cellulose hydrogel with microgroove-like structures. (a) Bright field image showing NRVMs cultured on the hydrogels for 5 days. (b-d) Confocal image and corresponding magnified images show sarcomeric α-actinin (SAA) in red and nuclei (DAPI) in blue. SAA staining at high magnification on showing the unidirectional growth of NRVM on both bridge (c) and valley (d) region of the hydrogel and a clear sarcomere structure in the up-right picture of d. (e) Mitochondria (MitoTracker in Green) enriched in NRVMs on both bridge region and valley region of ACH. (f) Confocal z-stacks scanned and rebuilt in 3D by ImageJ to show cell contributions in the cross-sections (a’, b’, c’, the red color in the heat map indicates cells, while the blue color indicates the microgroove-like hydrogel surface). (g, h) Bright field images of NRVMs cultured on the anisotropic hydrogels (g) and coverslips (h). The cell elongation directions are highlighted by the red and blue arrows on the anisotropic hydrogel and coverslip, respectively. (i) Quantitative analysis of the relative direction of NRVMs in both the experimental group (hydrogel; n=70) and the control group (coverslip; n=50). CONCLUSION In conclusion, diversified anisotropic hydrogels were fabricated successfully from cellulose solution in an alkali/urea aqueous system by combining of facile pre-stretching and acid treatment processes. The removable cellulose rigid chains and molecular bundles in the loosely chemical cross-linked gel rapidly rearranged along the pre-stretching direction, and then 20 ACS Paragon Plus Environment

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chemically and physically double cross-linked networks with oriented nanofibers formed through strong self-aggregation forces. Moreover, anisotropic hydrogel with a microgroove-like structure was obtained by releasing the pre-stretched elastic chemical networks, which could shrink upon removal of the external force, leading to the formation of parallel-distributed microgrooves along the perpendicular direction on the surface. The cellulose hydrogels exhibited a highly ordered nanofibrous architecture, anisotropic mechanical properties and optical birefringence. Importantly, the tensile strength and elastic modulus of the anisotropic cellulose hydrogels with high pre-stretching draw ratios in the parallel direction dramatically increased. The resulting hydrogel promoted well the arranged adhesion and orientation of cardiomyocytes, which showed potential physiological function in vitro. This work provides a new strategy for the facile fabrication of 3D-shaped anisotropic hydrogels with biocompatibility from natural polymer via the bottom-up method. ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI:xxxx. Hierarchical structure of wood; morphology of ACHs including ACH-1.4, ACH-1.8 and ACH2.2; water content of ACHs with various pre-stretching DRs; X-ray diffraction patterns of celluloses with various pre-stretching DRs; 2D WAXS data processing method, azimuthalintegrated intensity distribution curves, and Herman’s orientation parameter calculated from the corresponding curves; photographs showing the anisotropic mechanical property of ACH; culturing of NRVMs on ACHs without microgrooves; high resolution confocal image showing



the unidirectional growth of NRVMs with clear sarcomere strcuture on the surface of wrinkled cellulose hydrogel; photograh of mitochondria in NRVMs on the surface of ACH (PDF). Supplimentary Movie 1. Mechanical property of LCG (WMV). Supplimentary Movie 2. Anisotropic mechanical properties of cellulose hydrogel (WMV). Supplimentary Movie 3. Spontaneous beating of NRVMs cultured on anisotropic hydrogel with microgroove-like structure (WMV). AUTHOR INFORMATION Corresponding Authors *[email protected] *[email protected] OCRID Dongdong Ye: 000-0002-3377-0656 Donghui Zhang: 0000-0002-8128-5874 Liangbin Li: 0000-0002-1887-9856 Pingchuan Sun: 0000-0002-5603-6462 Chunyu Chang: 0000-0002-3531-5964 Lina Zhang: 0000-0003-3890-8690 Notes The authors declare no competing financial interest.


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Robust Anisotropic Cellulose Hydrogels Fabricated via Strong Self

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