Creating Biomimetic Anisotropic Architectures with Co-Aligned

Publication Date (Web): May 30, 2018 ... However, it is still an extreme challenge to create a three-dimensional (3D) scaffold with both aligned nanof...
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Creating Biomimetic Anisotropic Architectures with Co-Aligned Nanofibers and Macrochannels by Manipulating Ice Crystallization Linpeng Fan, Jing-Liang Li,* Zengxiao Cai, and Xungai Wang* Institute for Frontier Materials, Deakin University, Geelong, Victoria 3216, Australia S Supporting Information *

ABSTRACT: The continuous evolution of tissue engineering scaffolds has been driven by the desire to recapitulate structural features and functions of the natural extracellular matrix (ECM). However, it is still an extreme challenge to create a three-dimensional (3D) scaffold with both aligned nanofibers and aligned interconnected macrochannels to mimic the ECM of anisotropic tissues. Here, we develop a facile strategy to create such a scaffold composed of oriented nanofibers and interconnected macrochannels in the same direction, with various natural polymers typically used for tissue regeneration. The orientation of nanofibers and interconnected macrochannels can be easily tuned by manipulating ice crystallization. The scaffold demonstrates both structural and functional features similar to the natural ECM of anisotropic tissues. Taking silk fibroin as an example, the scaffold with radially oriented nanofibers and interconnected macrochannels is more efficient for capturing cells and promoting the growth of both nonadherent embryonic dorsal root ganglion neurons (DRGs) and adherent human umbilical vein endothelial cells (HUVECs) compared to the widely used scaffold types. Interestingly, DRGs and neurites on the SF scaffold demonstrate a 3D growth mode similar to that of natural nerve tissues. Furthermore, the coaligned nanofibers and macrochannels of the scaffold can direct HUVECs to assemble into blood vessel-like structures and their collagen deposition in their arrangement direction. The strategy could inspire the design and development of multifunctional 3D scaffolds with desirable structural features for engineering different tissues. KEYWORDS: anisotropic 3D scaffolds, aligned nanofibers, aligned macrochannels and macropores, silk fibroin nanofibers, alginate nanofibers, gelatin nanofibers, neurites, vascularization

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aligned nanofibers play a key role in supporting movement and mechanical load.11,14 Apart from aligned nanofibers, in anisotropic tissues such as nerve tissues, interconnecting macrochannels along the long axis of tissues provide space for the oriented growth of nerve fibers and facilitate the exchange and transport of oxygen, nutrients, and waste.15−17 Different techniques, especially electrospinning,2,18 3D printing,19,20 and lyophilization,15,21,22 have been widely used to produce scaffolds for tissue engineering applications. However, none of them has been capable of coupling aligned nanofibers with aligned macrochannels which always go in the opposite direction. Although electrospinning is able to produce 2D aligned nanofibrous films, the films have very small pores and low porosity due to the mechanical stretching or highspeed drawing during fabrication. As a result, cells cannot infiltrate into the films and grow only on their surface.23−25

caffolds formed by natural polymers such as proteins and polysaccharides play crucial roles in tissue engineering due to their excellent biocompatibility and biodegradability.1−4 An ideal scaffold should not only provide a threedimensional (3D) environment and support but also direct cell behaviors and functions by interacting with cells and mediating the complex multicellular interactions spatially and temporally.5−7 To this end, it is important for a scaffold to mimic the structural features and functions of the natural extracellular matrix (ECM). A natural ECM has a 3D porous and intricate architecture of collagen nanofibers with a diameter between 50 and 500 nm.8 In many tissues with anisotropic structural characteristics (such as dural, tendon, ligament, tympanic, and muscle tissues), cells and ECM nanofibers are highly aligned.9−14 These specific alignments support the specific physiological functions of tissues and organs. For example, radially aligned nanofibrous matrices of the dural and tympanic tissues are essential for carrying blood and conducting sound, respectively.9,13 In skeletal muscle, tendon, and ligament tissues, longitudinally © XXXX American Chemical Society

Received: March 3, 2018 Accepted: May 30, 2018 Published: May 30, 2018 A

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Scheme 1. Fabricating 3D Scaffolds with Radially Co-Aligned Nanofibers and Interconnected Macrochannels (A(F&C) Scaffolds) by a Facilely Guided Ice-Crystal Growth and Nanofiber Assembly Strategy, with Silk Fibroin (SF) as an Examplea

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The bottom pictures in (b) and (d) are digital photographs of the AF scaffold and the A(F&C) scaffold, respectively.

Meanwhile, these 2D films cannot mimic the extracellular matrix of native anisotropic tissues to provide support to cells and tissues in a 3D space.26−28 Three-dimensional printing is an emerging technique that offers greater flexibility in the tailored design of scaffolds, whereas the resolution of the printed structures is generally on micrometer or submillimeter scales.19,20 Creating hierarchical nanostructures presents an enormous challenge to this technique. Lyophilization is widely used to fabricate 3D porous scaffolds. Typically, a solution of polymer is frozen in a freezer. Upon sublimation of ice crystals, a porous polymer scaffold forms. However, anisotropic scaffolds with oriented nanofibers and macrochannels also have not been achieved with this technique to date. This is due to uncontrolled ice nucleation and growth, which leads to the formation of random pores and structures. The pore characteristics of a scaffold depend on how ice nucleates and grows in the polymer matrix. The size and morphology of ice crystals in turn determine the structure of the solid part of a scaffold. Neglecting the interplays between these two components, i.e., polymer and ice, is the reason for failing to fabricate scaffolds with all of the structural features. Furthermore, the interconnectivity of the pores in the scaffolds, which generally have a wall-like (nonfibrous) structure, is not sufficient for the infiltration, migration, and growth of cells and tissues.15,29 It also limits the transport of oxygen, nutrients, and wastes, which is a general cause of cell necrosis and tissue growth failure.15,30,31 In this work, we show that 3D scaffolds with both aligned nanofibers and aligned interconnected macrochannels can be created with various biomacromolecules, including silk fibroin (SF), using a facile guided ice-crystal growth and nanofiber assembly strategy. First, oriented fine ice crystals were created

to guide the assembly of oriented nanofibers. In brief, a solution of silk fibroin was immersed in a medium with a very low temperature (e.g., liquid nitrogen, Scheme 1a). Such a low temperature created a high temperature gradient in the solution, leading to the formation of radially aligned fine ice crystals, among which biomacromolecules were assembled into nanofibers (Scheme 1, top view of (a)). Following the removal of the fine ice crystals by sublimation (Scheme 1(i)), 3D scaffolds (AF, A denotes “aligned” and F denotes “nanofibers”) composed of radially aligned nanofibers were obtained (Scheme 1b). Second, the oriented nanofibers guided the oriented growth of large ice crystals along the fiber direction into the scaffold, which in turn led to further assembly of the nanofibers to form aligned macrochannels in their long-axis direction. In short, after treatment in ethanol to fix the protein structure of the nanofibers (to make them insoluble in water) (Scheme 1(ii)), the AF scaffold with radially aligned nanofibers was immersed in water and frozen at −20 °C (Scheme 1c). In this step, the relatively low temperature gradient and cooling rate led to the formation of large ice crystals. The radially aligned nanofibers of the AF scaffold guided the growth of large ice crystals into the scaffold in the alignment direction of the nanofibers (Scheme 1c and its top view). The removal of the large ice crystals by lyophilization led to the formation of macrochannels coaligned with the nanofibers, where the aligned nanofibers assembled to form walls of the macrochannels (Scheme 1d and its top view). Because of squeezing and the further assembly of nanofibers during macrochannel formation, pores on the nanofibrous channel walls were created, causing the macrochannels to be interconnected (Scheme 1d, top view). Meanwhile, a central macrochannel from the top to the bottom of the scaffold was formed to interconnect with all B

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Figure 1. Three-dimensionally aligned nanofibrous scaffolds with various geometries, diameters, and thicknesses as well as different fiber orientations. (a) Representative SEM images of 3D-aligned silk fibroin (SF) nanofibrous scaffolds (AF scaffolds in Scheme 1b). The fast Fourier transform (FFT) pattern (inset on the bottom left of the AF image in (a)) suggests that the nanofibers are well aligned along the radial direction of cylinder scaffolds (the red arrows indicate the alignment direction). Scale bars: 2, 1, and 10 μm from left to right, respectively. (b) Directionally freezing SF solutions in liquid nitrogen by self-made devices produces 3D SF nanofibrous scaffolds with controllable nanofiber orientations indicated by red arrows as well as various geometries (including cylinders, tubes, and particles), diameters, and thicknesses.

microscopically there is always a concentration heterogeneity. The nucleation of nanoparticles is thermodynamically favored at the locations where the concentration of SF is higher. However, since the overall concentration of free SF after nanofiber formation is low, isolated nanoparticles rather than continuous structures (such as nanofibers) formed on the existing nanofibers. Quenching an SF solution is facile and allows the fabrication of samples with various geometries (such as cylinders, tubes, and particles), diameters, and thicknesses (Figure 1b). In addition, the orientation of nanofibers can be easily varied by controlling the direction of the temperature gradient. For example, apart from cylinders and tubes with radially aligned nanofibers, cylinders and tubes with vertically aligned nanofibers can also be fabricated by slowly lowering the SF solution-containing tube into liquid nitrogen to induce a vertical temperature gradient. (The red arrows indicate the alignment direction of nanofibers; Figure 1b(i),(ii).) Furthermore, by directly dropping SF solution into liquid nitrogen, particles with nanofibers radially aligned toward the center of the particle were obtained (Figure 1b(iii)). We observed that fast freezing with a high temperature gradient is beneficial to the formation of nanofibers and directional structures. Instead of using liquid nitrogen, SF solutions contained in the same glass tubes were frozen in freezers at −80 and −20 °C, respectively, followed by removing ice crystals with a freeze drier (i.e., the conventional lyophilization method). SF scaffolds from freezing at −80 °C have a hybrid structure consisting of random short channels/ pores/nanofibers, but the short channels and pores are not interconnected (Supporting Information, Figure S1a). In the following study, the hybrid 3D SF scaffold from freezing at −80 °C is presented as W&F, where W denotes walls of the channels and pores and F denotes nanofibers. In comparison, only random pores were observed in the SF scaffolds from freezing at −20 °C, and the pores are also not well connected to form a network (Supporting Information, Figure S1b).

radially aligned macrochannels (Scheme 1d). The scaffold with radially coaligned nanofibers and macrochannels was indicated as A(F&C), where A represents “aligned”, F represents “nanofibers”, and C represents “channels”. Effects of the resulting silk fibroin (SF) scaffold on cell behavior and fate were investigated using the classical nonadherent embryonic dorsal root ganglion neurons (DRGs) and adherent human umbilical vein endothelial cells (HUVECs), respectively. In comparison to traditional 3D porous SF scaffolds, our 3D SF scaffolds with coaligned nanofibers and interconnected macrochannels were more efficient in capturing both adherent cells and nonadherent cells. Furthermore, the scaffold not only significantly promoted cell proliferation but also promoted the 3D growth of DRGs and neurites, which is similar to the growth mode of natural nerve tissues. More interestingly, HUVECs in the scaffold were directed to assemble into blood vessel-like structures and have their collagen deposition in the alignment direction of nanofibers and macrochannels.

RESULTS AND DISCUSSION Formation, Structure, and Features of Three-Dimensional Scaffolds. Figure 1a demonstrates the nanofibrous structure of the AF scaffold with radially aligned nanofibers, obtained after the removal of the radially aligned fine ice crystals created during directional freezing of an SF solution in liquid nitrogen. The nanofibers have a smooth morphology with an average diameter of 276 ± 78 nm. The fast Fourier transform (FFT) pattern (inset on the bottom left of the AF image in Figure 1a) shows that the nanofibers are well aligned along the radial direction of cylinder scaffolds. (The red arrows indicate the alignment direction of scaffold nanofibers.) Interestingly, the nanofibers are decorated with nanoparticles. The formation of nanoparticles could be due to the further nucleation of free SF molecules during the process of freezedrying for ice sublimation. In a solution of macromolecules, C

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Figure 2. Hierarchical structure of 3D SF scaffolds. (a) Micro-CT images of aligned macrochannels in the A(F&C) SF scaffold. Scale bars: 1000 μm. White arrows indicate the direction of radially aligned macrochannels in cylinder scaffolds. (b) SEM images of the macrochannel wall of the A(F&C) SF scaffold at various magnifications, revealing the aligned SF nanofibers decorated with nanoparticles and pores along the long-axis direction of the aligned macrochannel. Yellow arrows indicate the direction of radially aligned nanofibers along the long-axis direction of the aligned macrochannel. The green, blue, and red arrows indicate the aligned nanofiber, the nanoparticle, and the pore, respectively. Scale bars: 10, 2, and 1 μm from left to right, respectively. (c) Schematic dimension presentation of the relevant structures of the A(F&C) scaffold. (d) Micro-CT and SEM images of the W&F scaffold showing a hybrid structure of random short channels/pores/nanofibers and the W scaffold revealing a wall-like structure. Scale bars: 1000 and 100 μm for Micro-CT and SEM images, respectively.

Hence, when the temperature gradient and cooling rate are not high enough, large ice crystals generally form, facilitating the formation of large but not interconnected pores, and the scaffold has a wall-like structure. In the following study, the porous wall-like 3D scaffold from freezing at −20 °C is represented by W, which denotes wall-like structures. As described in Scheme 1, the target 3D A(F&C) scaffold with radially coaligned nanofibers and macrochannels was obtained after the removal of the radially aligned large ice

crystals in the 3D AF scaffold (Scheme 1c,d). Threedimensional micro-CT images (Figure 2a) and the movie (Supporting Information, Movie S1) demonstrate that each radially aligned macrochannel (diameter 100−1000 μm) connects the side surface and center of the A(F&C) scaffold. (SEM profiles of macrochannels in the A(F&C) scaffold were presented in Supporting Information, Figure S2.) As shown in Figure 2b, the channel walls are composed of SF nanoparticles and nanofibers (diameter 50−600 nm) that are well aligned D

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Figure 3. Mechanical characteristics of scaffolds and representative scaffold morphologies after compression. (a) Compressive modulus of scaffolds and their corresponding morphologies after the mechanical test: porous wall-like 3D silk fibroin (SF) scaffolds (W), 3D SF scaffolds with short channels/pores/nanofibers (W&F), and 3D SF scaffolds with radially coaligned nanofibers and macrochannels (A(F&C)). (b) Stress−strain plots of 3D SF scaffolds. (c) Hysteresis loops during the first loading cycle for 3D SF scaffolds. (d) Peak stresses of 3D SF scaffolds during the first loading cycle.

along the channel direction (indicated by yellow arrows). Many pores with diameters in the range of 50−1000 nm can be identified on the channel wall, and these pores cause the channels to be interconnected. The pores are also aligned along the same direction as nanofibers. The formation of pores could be due to the contraction of nanofibers during the drying process. More interestingly, a central channel (diameter 0.4−2 mm) from the top to the bottom of the scaffold was created spontaneously (Scheme 1d). This special structure could be produced by the contraction of the scaffold during the lyophilization for ice sublimation. All of the relevant sizes of the structures within the hierarchical 3D A(F&C) scaffold are summarized in Figure 2c. The macrochannels with porous nanofibrous walls in the A(F&C) scaffold and the central channel are very important for cell and tissue infiltration as well as their growth by providing space and faciliating the transport of oxygen, nutrients, and wastes. The aligned nanofibers on channel walls can play an important role in promoting cell capture and proliferation as well as directing cell migration and mediating the complex multicellular interactions. Furthermore,

nanofibers and nanoparticles can be good carriers for the delivery of growth factors or drugs. The A(F&C) scaffolds have also been achieved from mixtures of SF and gelatin as well as other biomacromolecules such as sodium alginate (Supporting Information, Figure S3). Without the guidance of the aligned nanofibers, macrochannels cannot be made in randomly porous W and W&F scaffolds under the same conditions (Figure 2d; Supporting Information, Figure S4 and S5; Supporting Information, Movies S2 and S3 for W&F and W scaffolds, respectively). The random stiff wall-like structures in the W and W&F scaffolds caused the growth of ice crystals to be confined to the free space of the scaffolds, and thus no long channels formed. Secondary Structure and Mechanical Characteristics of Three-Dimensional Scaffolds. To make the scaffolds insoluble in water, an ethanol post-treatment (Scheme 1(ii)) was used to transform the random coil structure of SF to the βsheet crystalline structure. Silk fibroin is water-insoluble when β-sheets are dominant.18,32 The secondary-structure change in SF can be indicated by the shift of its characteristic absorption E

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Figure 4. Co-aligned nanofibers and macrochannels of 3D A(F&C) scaffolds facilitate the capture of embryonic dorsal root ganglion neuron cells (DRGs) and direct the 3D growth of DRG neurites. (a) Viability (MTS absorbance index) of DRGs captured by radially aligned 3D SF nanofibrous scaffolds without channels (AF), porous wall-like 3D SF scaffolds (W), 3D SF scaffolds with short channels/pores/nanofibers (W&F), and 3D SF scaffolds with radially coaligned nanofibers and macrochannels (A(F&C)). (b) Confocal fluorescence microscopy images reveal that the structures of W, W&F, and AF scaffolds limited DRGs and DRG neurites to grow on the surface of the scaffolds only. Scale bars: 100 μm for the W and W&F scaffolds and 50 μm for the AF scaffold. (c) Co-aligned nanofiber- and macrochannel-directed 3D growth of DRG neurites in the 3D A(F&C) scaffold. Scale bars: 75, 25, and 25 μm from left to right, respectively.

peaks (1600−1500 cm−1 for amide II and 1700−1600 cm−1 for amide I) in ATR-FTIR spectra.18,32,33 Before post-treatment with ethanol, all three types of SF scaffolds showed one of the main characteristic peaks at around 1644 cm−1 (which suggests random coils) (Supporting Information, Figure S6a).18 It is noteworthy that the scaffolds that formed at −20 and −80 °C (W and W&F, respectively) presented another main characteristic peak at 1517 cm−1, which indicates dominant β-sheets.18 However, the scaffold (AF) formed by freezing in liquid nitrogen (about −196 °C) showed another main characteristic peak at 1533 cm−1, which suggests dominant random coils.18 These results suggest that the freezing treatment in liquid nitrogen could be beneficial to the formation of random coils in SF scaffolds (Supporting Information, Figure S6a). After posttreatment with ethanol, all three types of SF scaffolds presented main characteristic peaks at around 1700, 1622, and 1517 cm−1, suggesting that the ethanol-treated scaffolds mainly consist of β-sheets (Supporting Information, Figure S6b).18,32 Mechanical characteristics of different scaffolds were given in Figure 3. The A(F&C) scaffold (with coaligned nanofibers and

macrochannels) has a compressive modulus of around 80 kPa, which is lower than that of the porous wall-like scaffold (W) and the scaffold with short channels/pores/nanofibers (W&F) (around 100 and 140 kPa, respectively) (Figure 3a). The lower modulus of the A(F&C) scaffold could be due to its large channel-based structure with nanofibers. It is noteworthy that the A(F&C) scaffold did not fracture even if it was compressed to approximately 85% of its original height (Figure 3b). Furthermore, after being compressed in the mechanical test, the A(F&C) scaffold maintained radially well aligned morphology and structure, with just some minor collapses seen on its surface that are probably due to damage to some macrochannels (Figure 3a). Although the A(F&C) scaffold has a relatively low peak stress (Figure 3d), its stress−stain curve under cyclic compression shows a hysteresis loop (Figure 3c), confirming that its viscoelastic nature is similar to that of the natural ECMs.34−36 Scaffolds with Co-aligned Nanofibers and Macrochannels Facilitating DRG Capturing and the ThreeDimensional Growth of Neurites. To understand the effects F

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Figure 5. Three-dimensional A(F&C) scaffolds with radially coaligned nanofibers and macrochannels enhance the capture and proliferation of adherent human umbilical vein endothelial cells (HUVECs) and direct cell migration and growth. (a) Viability (MTS absorbance index) of HUVECs captured by radially aligned 3D silk fibroin (SF) nanofibrous scaffolds without macrochannels (AF), porous wall-like 3D SF scaffolds (W), 3D SF scaffolds with short channels/pores/nanofibers (W&F), and 3D SF scaffolds with radially coaligned nanofibers and macrochannels (A(F&C)). (b) Viability (MTS absorbance index) of HUVECs in 3D AF, W, W&F, and A(F&C) scaffolds after different periods of culture. (c) Scheme illustrating how to read the images presented in (d). (d) Growth of HUVECs in 3D AF, W, W&F, and A(F&C) scaffolds after 3 days of culture. All of the scaffolds were stained by two fluorescent dyes. To clearly show the morphology of cells and scaffolds, a blue color was used for the W and W&F scaffolds and a red color was used for the AF and A(F&C) scaffolds. Scale bars: 25 μm in W, W&F, AF, and inset 1; 75 μm in A(F&C).

of the coaligned nanofibers and macrochannels on cells, the ability of the scaffolds to capture cells and promote their growth was investigated using embryonic dorsal root ganglion neurons (DRGs). As shown in Figure 4a, the A(F&C) scaffold (with coaligned nanofibers and macrochannels) demonstrated superior DRG capture capacity. The AF scaffold (with aligned nanofibers but without channels) showed the lowest DRG capture. Figure 4b illustrates the areas of scaffolds that were scanned and the corresponding images. Obviously, affluent neurites were aligned in the direction of nanofibers on the surface of the AF scaffold, but they were not clearly observed in the inner of this scaffold, as shown by the image of the AF cross-section. In the porous W and W&F scaffolds, neurites were also mainly aggregated on the surface. In the interior of the porous wall-like W scaffold, the neurite infiltration of aggregated DRGs occurred along the pore walls only (Supporting Information, Figure S7). In the interior of the W&F scaffold (with short channels/pores/nanofibers), the pore walls also led to the aggregation of DRGs and limited the neurite outgrowth (Supporting Information, Figure S7). These results indicate that in the absence of the natural nerve conduit-

like channels it is difficult for DGRs to grow in the interior of these scaffolds during 21 days of culture, and thus the neurite outgrowth and extension of DRGs were suppressed. In comparison to the structures of the scaffold AF, W, and W&F, the radially aligned macrochannels toward the center of scaffolds provided enough space similar to the natural nerve conduit for migration and the 3D growth of the DRGs and neurites. Figure 4c illustrates the scanned areas of the A(F&C) scaffold and the corresponding images. In the A(F&C) scaffold, DRGs can be clearly seen, and many long neurites had grown through the macrochannels. (The channels, channel walls, and neurites were indicated by white arrows.) Interestingly, a closeup view of the channels revealed that DRGs and neurites mainly grew along the channels, suggesting a 3D growth mode of neurites similar to that of natural nerve fibers.17 This is totally different from the 2D growth of DRGs and neurites along the aligned nanofibers on the surface of the AF scaffold (Figure 4b). From the last image (inside of A(F&C)) in Figure 4c, neurites in bundles are obvious, which is very important for the formation of nerve tissues.17 These observations demonstrate that the coaligned nanofibers and macrochannels can not G

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Figure 6. Co-aligned nanofibers and macrochannels in 3D scaffolds direct the formation of the CD31-positive vessel-like structures and the collagen deposition of HUVECs by regulating the growth, migration, and interaction of cells. (Figure 5c illustrates how to read the images presented here.) (a) Growth and interaction of HUVECs in the 3D silk fibroin (SF) scaffold with radially coaligned nanofibers and macrochannels (A(F&C)), the radially aligned 3D SF nanofibrous scaffold without channels (AF), the 3D SF scaffold with short channels/ pores/nanofibers (W&F), and the porous wall-like 3D SF scaffold (W). Scale bars: 50 μm in A(F&C), W&F, and W; 25 μm in AF. (b) Sequential confocal slices of the A(F&C) channel shown in (a) demonstrating many vessel-like structures formed in the long-axis direction of the channel. Scale bars: 50 μm. (c) Collagen deposition of HUVECs in the A (F&C), AF, W&F, and W scaffolds. For the AF scaffold, the deposited collagen only on the surface of this scaffold was stained and imaged. (It was difficult to observe the deposited collagen inside this scaffold due to a small number of cells.) The blue arrows indicate the deposited collagen stained in bright red. The black arrow in AF indicates the direction of aligned nanofibers, and the black arrow in A(F&C) indicates the direction of the channel. Scale bars: 100 μm in W, W&F, and AF; 200 μm in A(F&C).

only promote the adhesion of nonadherent DRGs but also direct them to grow, migrate, and interact in the 3D space similar to that of the natural ECM. Scaffolds with Co-aligned Nanofibers and Macrochannels Promoting HUVECs Capturing and Growth. Effects of the coaligned nanofibers and macrochannels on cells were further confirmed using adherent human umbilical vein endothelial cells (HUVECs). At all time points, the A(F&C) scaffold demonstrated a significantly higher capacity of cell capture and proliferation than did the porous wall-like 3D silk fibroin (SF) scaffold (W) and the 3D SF scaffold with short channels/pores/nanofibers (W&F) (Figure 5a,b). This indicates that the aligned macrochannel and nanofibrous structures of the A(F&C) scaffold are beneficial to cell capture and proliferation. Compared to the W scaffold, the W&F scaffold demonstrated higher cell adhesion and proliferation

viabilities after 8 h and 6 days, respectively, which is probably due to the presence of nanofibers in the W&F scaffold. These findings are consistent with the observations from other researchers that nanofibers can promote cell adhesion and proliferation, probably by providing more cues and binding sites.5 To further identify the effect of macrochannels, the AF scaffold (with radially aligned nanofibers, but without channels) was also used as a cell culture substrate in this study (Figure 5a,b). Obviously, the A(F&C) scaffold demonstrated significantly higher cell viability than the AF scaffold at all time points, confirming the advantages of macrochannels in cell capture and proliferation. Furthermore, the W and W&F scaffolds also had higher cell viabilities in comparison to the AF scaffold. This is probably due to the fact that the W and W&F scaffolds have more space for cell adhesion and proliferation with their larger pores. H

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Information, Figure S8.) A comparison between Figures 6 and 4 indicates that the adherent HUVECs were mainly directed by the aligned nanofibers on the channel wall of the A(F&C) scaffold, and the nonadherent DRGs and neurites preferred to grow along the 3D channel space. Fourteen sequential confocal slices of the A(F&C) macrochannel in Figure 6a are presented in Figure 6b. Clearly, there are many vessel-like structures aligned on the wall of the channel in the interior of the A(F&C) scaffold. These findings demonstrate that the coaligned nanofibers and macrochannels facilitated the spreading, migration, elongation, and interaction of HUVECs to assemble into vessel-like structures. More interestingly, the structure of the A(F&C) scaffold not only directed the behavior of cells but also directed the collagen deposition of cells in the scaffold. As shown in Figure 6c, in the W and W&F scaffolds, the collagen deposition of HUVECs was in a random arrangement. (After staining, the deposited collagen was bright red, as indicated by the blue arrows in Figure 6c.) For the AF scaffold, the deposited collagen only on the surface of the scaffold was stained and imaged. (It was difficult to observe the deposited collagen in the inside of this scaffold due to the lack of cells.) Although collagen was deposited along the nanofiber direction (indicated by the black arrow in Figure 6c) of the AF scaffold, the matrix was not well aligned, which may be due to the flat and polygonous arrangement of HUVECs on the loosely aligned nanofibers, as shown in Figures 5d and 6a. Obviously, in the A(F&C) scaffold, the collagen was well deposited along the aligned nanofibers of channel walls, which is consistent with the good alignment and elongation behaviors of HUVECs on the channel wall of this scaffold, as indicated in Figures 5d and 6a,b. Stability of Three-Dimensional Scaffolds. In order to investigate the in vitro degradation of 3D SF scaffolds, the scaffolds were incubated in PBS and protease/PBS solution for 1, 5, 10, 15, 21, and 28 days (Supporting Information, Figure S9a). After being incubated in PBS for up to 28 days, all three types of scaffolds experienced only a small loss in weight (≤5%). In the presence of protease, the weight loss of the A(F&C) scaffold is about 54%, while those of W and W&F scaffolds are around 38 and 33%, respectively. The higher weight loss of the A(F&C) scaffold at all time points may be due to the fact that its nanofibrous structure provides a greater surface area to interact with protease molecules. To provide further insight into the degradation of the scaffolds, the morphologies of the scaffolds after incubation in the solutions for 10 days were examined using SEM (Supporting Information, Figure S9b and S10). In the presence of protease, some small pores (indicated by red arrows) on the surface of nanofibers or walls of the scaffolds were observed. Meanwhile, some cracks on the relatively thin nanofibers in the A(F&C) scaffold were seen, as indicated by the blue arrows in Figure S9b (Supporting Information). Despite these, all three types of scaffolds maintained good morphology and integrity even in the presence of protease, as shown by SEM images taken at low magnification (Supporting Information, Figures S9b and S10). The excellent stability of the scaffolds was also demonstrated in the in vitro cell-culture studies in which the channels still showed good morphology and structure after 21 days of cell culture (Figures 4c, 5d, and 6a,b). With the pure PBS solution, no obvious changes in the surface of the nanofibers of the A(F&C) scaffold and the walls of the W and W&F scaffolds were identified. These results suggest that although the

To gain more insight into the effects of the coaligned nanofibers and macrochannels, cells grown in the scaffolds for 3 days were imaged using confocal fluorescence microscopy (Figure 5d). To date, it remains a problem that cell behaviors including cell spreading, migration, elongation, and interaction are often hindered due to the small pores and their low interconnectivity of scaffolds as well as the absence of binding and guiding cues in a scaffold.15,29 This is also true for both the W and W&F scaffolds. As shown in Figure 5d, cell spreading was significantly limited by pore walls (indicated by yellow arrows in (W)) or presented with blunt edges (indicated by white arrows in (W&F)) as if cells were cultured on the surface of a flat material. Although cells were also observed in the AF scaffold, it was difficult to find them during scanning under confocal microscopy due to the small number of cells inside this scaffold. Cells in the AF scaffold were not well aligned and elongated in the direction of nanofibers, exhibiting a relatively flat and polygonous morphology. This is probably due to the fact that the loosely aligned nanofibers provided cells with many surrounding signals from different directions.5 Obviously, cells on the channel wall of the A(F&C) scaffold were well elongated and aligned along with nanofibers, and they seemed to be in their dynamic migrating modes. The presence of large 3D channels reduced the space in the scaffold so that nanofibers were compacted on the walls of the channels, providing cells with more signals in the long-axis direction of nanofibers. (The directions of channels and nanofibers were indicated by white arrows.) This could explain the cell growth and morphologies observed in the A(F&C) scaffold. Scaffolds with Co-aligned Nanofibers and Macrochannels Directing Vascularization and Collagen Deposition. The proliferation, migration, and interaction of endothelial cells are very important for the formation of the tubal structures in both vasculogenesis and angiogenesis.37 HUVECs are a classic endothelial cell model for studying vascularization.38,39 As observed above (Figure 5), the A(F&C) scaffold can promote the proliferation of HUVECs. Hence, it was anticipated that the cell migration and elongation induced by the coaligned nanofibers and macrochannels would enhance intercellular interactions to facilitate the formation of vessel-like structures. To prove this, we cultured HUVECs for up to 21 days to observe vascularization behaviors of cells in the different scaffolds (Figure 6a,b; Figure 5c illustrates how to read the images in Figure 6). Clearly, all cells are CD31-positive (CD31 is a glycoprotein expressed on endothelial cells), where the CD31-positive cells are in bright green, with their cell nuclei in bright blue (Figure 6a,b and Supporting Information, Figure S8). This suggests they maintained the characteristics of HUVECs in the scaffolds after long-term culture. In the W and W&F scaffolds, many cells exhibited round morphology with just a few nuclei elongated (Figure 6a,b and Supporting Information, Figure S8). Obviously, the spreading, migration, and elongation of cells were limited by scaffold walls, leading to the local aggregation and interaction of some cells. In the AF scaffold, although some cell nuclei were elongated, most of the cells were not significantly aligned and elongated, presenting a polygonous morphology (Figure 6a,b and Supporting Information, Figure S8). Interestingly, in the A(F&C) scaffold, all cells and cell nuclei were elongated and aligned on the wall of the channels where they interacted and assembled into CD31positive vessel-like structures (The channel, channel walls, vessel-like structures, and aligned and elongated cell nuclei are indicated by white arrows; Figure 6a,b and Supporting I

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ACS Nano

ASSOCIATED CONTENT

scaffolds are biodegradable, their morphology and integrity can be maintained for a long time.

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.8b01648. Experimental methods and results (PDF) A(F&C) scaffold (MPG) W&F scaffold (MPG) W scaffold (MPG)

CONCLUSIONS We have developed a facile strategy for guiding the growth of ice crystals and the assembly of nanofibers to create a biomimetic anisotropic 3D scaffold (the A(F&C)) with coaligned nanofibers and macrochannels using various natural polymers such as silk fibroin (SF). As a model platform for cell culture and study in vitro, the 3D SF A(F&C) scaffold showed significantly higher cell capture and growth-promoting capability than the widely used porous 3D SF scaffolds and the 3D aligned SF nanofibrous scaffold without macrochannels for both nonadherent DRGs and adherent HUVECs. More importantly, the coaligned nanofibers and macrochannels of the A(F&C) scaffold can not only direct the neurite growth of DRGs in the 3D space similar to the natural nerve conduit but also regulate the growth, migration, alignment, elongation,and interaction of HUVECs to assemble into blood vessel-like structures in the scaffold in vitro. It is interesting that the adherent HUVECs were mainly directed by the aligned nanofibers on the wall of the A(F&C) scaffold and the nonadherent DRGs and neurites preferred to grow along the 3D space of macrochannels. More interestingly, the HUVECs were also directed to have their collagen deposition along the coaligned nanofibers and macrochannels. In brief, the scaffold developed in this work served as an excellent model platform for the proof of concept that the creation of the ECMmimicking 3D structure is very important for providing insight into cell behaviors and functions. Considering the facile fabrication technology, the discovery in this work will provide inspiration for developing biomimetic functional 3D scaffolds based on aligned nanofibers and macrochannels for use in tissue engineering. For example, the tube scaffold with radially coaligned nanofibers and macrochannels could be beneficial to multilayer cell seeding for constructing blood vessel tissues. Likewise, with structure similar to that of the natural nerve conduit, the column scaffold containing coaligned nanofibers and macrochannels in the scaffold long-axis direction could provide a better support for nerve regeneration than the widely used hollow tube scaffolds with thin walls.

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Jing-Liang Li: 0000-0003-0709-2246 Author Contributions

L.F., J.-L.L., and X.W. conceived the project. L.F., Z.C., J.-L.L., and X.W. designed the experiments. L.F. and Z.C. performed the experiments. L.F. wrote the draft of manuscript. All authors contributed to the analysis and discussions of the data and the revision of the manuscript. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS X.W. and J.-L.L. acknowledge the Australian Research Council (ARC) for support through an Industry Transformation Research Hub (IH140100018). J.-L.L. also acknowledges the ARC for support through a Future Fellowship project (FT130100057). REFERENCES (1) Sayyar, S.; Murray, E.; Thompson, B.; Chung, J.; Officer, D. L.; Gambhir, S.; Spinks, G. M.; Wallace, G. G. Processable Conducting Graphene/Chitosan Hydrogels for Tissue Engineering. J. Mater. Chem. B 2015, 3, 481−490. (2) Fan, L.; Cai, Z.; Geng, X.; Wang, H.; Li, J.; He, C.; Mo, X.; Wang, X. Fabrication and Characterization of Compound Vitamin B/Silk Fibroin Nanofibrous Matrices. J. Controlled Release 2017, 259, e85− e86. (3) Smith, B. D.; Grande, D. A. The Current State of Scaffolds for Musculoskeletal Regenerative Applications. Nat. Rev. Rheumatol. 2015, 11, 213−222. (4) Porter, D.; Vollrath, F. Silk as a Biomimetic Ideal for Structural Polymers. Adv. Mater. 2009, 21, 487−492. (5) Stevens, M. M.; George, J. H. Exploring and Engineering the Cell Surface Interface. Science 2005, 310, 1135−1138. (6) Lutolf, M.; Hubbell, J. Synthetic Biomaterials as Instructive Extracellular Microenvironments for Morphogenesis in Tissue Engineering. Nat. Biotechnol. 2005, 23, 47−55. (7) Richter, B.; Hahn, V.; Bertels, S.; Claus, T. K.; Wegener, M.; Delaittre, G.; Barner-Kowollik, C.; Bastmeyer, M. Guiding Cell Attachment in 3D Microscaffolds Selectively Functionalized with Two Distinct Adhesion Proteins. Adv. Mater. 2017, 29, 1604342. (8) Barnes, C. P.; Sell, S. A.; Boland, E. D.; Simpson, D. G.; Bowlin, G. L. Nanofiber Technology: Designing the Next Generation of Tissue Engineering Scaffolds. Adv. Drug Delivery Rev. 2007, 59, 1413−1433. (9) Xie, J.; MacEwan, M. R.; Ray, W. Z.; Liu, W.; Siewe, D. Y.; Xia, Y. Radially Aligned, Electrospun Nanofibers as Dural Substitutes for Wound Closure and Tissue Regeneration Applications. ACS Nano 2010, 4, 5027−5036. (10) Domingues, R.; Chiera, S.; Gershovich, P.; Motta, A.; Reis, R. L.; Gomes, M. E. Enhancing the Biomechanical Performance of

EXPERIMENTAL SECTION Silk Fibroin (SF) Solution Generation. SF solution (2%) was obtained by dissolving 2 g of a regenerated SF sponge (fabrication details in Supporting Information) in 100 mL of ultrapure water for further use. Three-Dimensional SF Scaffold Preparation. Scaffolds with Radially Aligned Nanofibers (AF). SF solution in a glass tube (12 mm diameter × 45 mm height) was directly immersed in liquid nitrogen. Radially aligned nanofibrous scaffolds were produced by removing ice crystals using a freeze-dryer. The fabrication scheme is shown in Scheme 1. To make the scaffolds insoluble in water, the scaffolds were treated by immersion in ethanol at ambient temperature for 12 h and then thorough rinsing with ultrapure water to obtain water-resistant AF scaffolds. Scaffolds with Coaligned Nanofibers and Macrochannels (A(F&C)). The AF scaffolds were immersed in ultrapure water and then frozen at −20 °C for 72 h. After removing ice crystals with a freeze-drier, A(F&C) scaffolds were obtained. J

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DOI: 10.1021/acsnano.8b01648 ACS Nano XXXX, XXX, XXX−XXX