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Bioinspired Multicompartmental Microfibers from Microfluidics Yao Cheng, Fuyin Zheng, Jie Lu, Luoran Shang, Zhuoying Xie, Yuanjin Zhao,* Yongping Chen,* and Zhongze Gu* Microfibers are long, thin, and flexible materials. They are useful for the creation of various functional three-dimensional (3D) objects, including clothes and other architectures by folding, bundling, reeling, and weaving.[1–6] These microfibers have also been attractive for creating complex 3D tissues in vitro because the hierarchical structures of the human body include various types of fiber-shaped 3D cellular constructions, such as blood vessels,[7–9] muscle fibers,[10] nerve bundles,[11] hepatic cord structures,[12] and similar anatomical structures.[13–15] However, the fiber-shaped structures in plants and animals usually possess compositional and topographical properties that vary spatiotemporally on the microscale.[16] Thus, to engineer these linear tissues, preparation of heterogeneous cellular microfibers with similar microstructures is essential for regulating the direction and distribution of cell growth.[17,18] Several methods have recently been developed to fabricate microfibers continuously, such as electrospinning, wet spinning, and melt spinning.[19–25] However, most of the microfibers generated by these techniques have a homogeneous chemical composition and structure, and the creation of continuous microfibers with spatiotemporally tunable chemical compositions and structures remains a great challenge. Microfluidics has created exciting avenues of scientific and engineering research into the synthesis of functional materials.[26–30] Microfluidics is the technology that deals with the precise control and manipulation of small quantities of fluids constrained in microchannels of small cross-sectional dimensions.[31] Because of this feature, microfluidics is considered to be a promising candidate for continuous fabrication of functional microfibers. A series of research projects has demonstrated the feasibility of engineering continuous microfibers with single or multiple compositions and structures by using coaxial polydimethylsiloxane (PDMS) microfluidic
Dr. Y. Cheng,[+] Dr. F. Y. Zheng,[+] Dr. J. Lu, Dr. L. R. Shang, Prof. Z. Y. Xie, Prof. Y. J. Zhao, Prof. Z. Z. Gu State Key Laboratory of Bioelectronics Southeast University Nanjing 210096, China E-mail: [email protected]; [email protected] Prof. Y. P. Chen School of Energy and Environment Southeast University Nanjing 210096, China E-mail: [email protected] [+]These authors contributed equally to this work
DOI: 10.1002/adma.201400798
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channels.[32–35] These systems have broken through several barriers associated with conventional methods.[36,37] However, due to the limitation of generating microchannels of fixed dimensions by current microfluidic techniques, it has not been possible to generate heterogeneous microfibers with complex morphologies and tunable compositions to the same extent as those present in natural organisms. Here, we present a one-step, continuous process for the scalable formation of microfibers with desired features using novel capillary microfluidics. Our method employs immediate microfiber gelation from multiflows of sodium alginate (Na-alginate) solution and calcium chloride (CaCl2) solution at the orifice of an injection capillary, which is coaxially aligned with a collection capillary inside a square capillary (Figure 1a–c). As these flows had a low Reynolds number, they mainly formed laminar flows in the microfluidic channels and only slowly diffusively mixed across the adjacent streams (Figure S1, detailed description in the Supporting Information). Thus, the resultant microfibers had the same structure as the injection flows. Homogeneous structures as well as microfibers possessing anisotropic multicompartmental body, core, and shell compositions could be generated by varying the injection capillary design. To our knowledge, such microfibers have not previously been reported. Furthermore, we have also explored the potential use of these heterotypic microfibers for tissue-engineering applications by creating multifunctional fibers with a spatially controlled encapsulation of cells. The proposed microfibers may enable the construction of fiber-shaped tissues in natural organisms. In a typical experiment, the inner phase of the Na-alginate precursor was pumped through the tapered capillary, while the CaCl2 stream was pumped in the same direction through the region between the inner capillary and the outer square capillary. Facilitated by hydrodynamic focusing, a 3D coaxial sheath flow stream around the flow of the precursor was formed at the merging of both flows. Thus, because of fast diffusioncontrolled ion cross-linking, a hydrogel microfiber was generated in situ. It is worth mentioning that by changing the flow rates, the microfibers could form four kinds of flow patterns, including clogging, wavy microfibers, straight microfibers, and spiral microfibers, in the collection channel (Figure S2, Supporting Information). Among these, the flow pattern of straight microfibers was desired (as is explained in the Supporting Information). Figure 1d and 1e show photographs of calcium alginate microfibers resulting from this. The microfibers had a cylindrical structure and high uniformity. The diameter and speed of generation of the microfibers was investigated. The speed can be increased by simply increasing the core
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Figure 1. a) Schematic illustration of the capillary coaxial microfluidic device used for continuously generating simple microfibers. b,c) Digital photographs of the microfluidic device and its microstructure during the formation of the microfibers. d) Digital image of continuous microfibers in a vessel. e) Microscope image of the alginate microfibers, the scale bar indicates 200 µm. f) Relationship between the diameter of the microfibers and the flow rates of the Na-alginate solution (blue color, with a fixed CaCl2 flow rate of 15 mL h−1) and calcium chloride solution (red color, with a fixed Na-alginate flow rate of 5 µL min−1); the orifice diameter of the tapered injection capillary of the microfluidic device is 50 µm.
and sheath flow rates. In a specific microfluidic device (with a tapered tip of 50 µm and a collection channel of 580 µm), the relationships between the microfiber diameter and the core and sheath flow rates are shown in Figure 1f. It was found that because of core-flow solidification, the practical diameters of the microfibers were larger than those theoretically calculated from the pure fluids (Figure S3, Supporting Information). The diameter of the microfibers increased in a flow-rate-dependent manner of the Na-alginate flow, showing an increase of about 90 µm as the sheath flow rate increased from 1 to 35 µL min−1. In contrast, as the CaCl2 sheath flow rate increased from 4 to 50 mL h−1, the fiber diameter decreased by 80 µm on average. A larger adjustment range of the microfiber diameter could be achieved by using different microfluidic devices (Figure S4, Supporting Information). To fabricate multicomponent microfibers, our basic microfluidic device underwent a series of transformations. As shown in Figure 2a, instead of the single-channel injection capillary previously described, a tapered two-barrel capillary was used as the injection channel and coaxially inserted into the collection capillary. Two fluids of Na-alginate solution mixed with different dyes were simultaneously pumped into the injection capillary channels using the same flow rate. Because of their low Reynolds number, they mainly formed a laminar flow and no obvious diffusive mixing occurred across the two color streams before being solidified (Figure 2b,c). When they were flowed into the collection channel and reacted with the CaCl2 sheath flow in situ, a two-compartment Janus calcium alginate microfiber formed stably, similar to the formation process of the single-compartment microfibers. Figure 2d represents typical cross-sectional confocal laser scanning microscopy (CLSM) images of the Janus microfibers. It was found that the internal architecture of the microfiber displayed an extremely sharp
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boundary at the interface between the two compartments, suggesting minimal mass transfer between the compartments. By replacing the two-barrel injection capillary with a four- or six-barrel injection capillary, four- and six-compartment calcium alginate fibers could correspondingly be generated, as shown in Figure 2e and 2f. Each compartment of the microfibers could have its own chemical composition, which is determined by the chemical composition of the initial injecting solutions. Thus, the multicompartmental microfibers have great potential applications in creating multiactive encapsulations. The volume ratio of each compartment could also be modulated by controlling the flow rates of the injection fluids. Hollow alginate microfibers were also fabricated using a microfluidics capillary with hierarchical injection channels, as indicated in Figure 3a. When three fluids, namely, a core fluid (CaCl2), a middle fluid (Na-alginate), and an outer sheath fluid (CaCl2), were introduced into the microfluidic device (Figure 3b), a hollow microfiber was generated in two stages. During the first stage, a coaxial flow of the core fluid and middle fluid was generated and the primary gelation process started along the inner wall of the hollow microfiber. In the second stage, the two coaxial fluids merged with the calcium ion sheath flow, and together they formed a three-layered coaxial flow. The gelation occurred simultaneously at both the inner and outer layers of the middle Na-alginate fluid. The three-layered coaxial flow moved through the collection capillary and the solidified alginate hollow microfiber was continuously generated and extruded (Figure 3c). The hollow microfibers achieved were of uniform shell thickness, as shown in Figure 3d and 3f. Both the diameter of the hollow microfibers and the thickness of the microfiber shell could be modulated by controlling the physical parameters of the fluids or tuning the flow rates of the fluids (as shown in Figure S5, Supporting
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COMMUNICATION Figure 2. a) Schematic illustration of the microfluidic device used for generating Janus-compartmental microfibers. b,c) Digital photographs of the microfluidic device and its microstructure during the formation of the Janus microfibers. d–f) Schematic illustrations of the multicompartmental microfibers and their corresponding microfluidic injection channels, and the cross-sectional CLSM images of the multicompartmental microfibers: d) Janus-compartments; e) four-compartments; f) six-compartments. The scale bar indicates 200 µm. The experimental flow rate conditions were QNa-alginate/QCaCl2 = 0.5:10; 0.3:10; 0.2:10 mL h−1, respectively in (d–f).
Information). Hollow microfibers with multiple core channels could also be generated by inserting several spindle capillaries in parallel to the original injection capillary for pumping of the core fluids (Figure 3g–i and Figure S6, Supporting Information). The hollow microstructures of these microfibers are confirmed by the scanning electron microscopy (SEM) images in Figure S7 in the Supporting Information.
To fabricate hollow microfibers with multiple shell layers, a microfluidic device was constructed with an increasing number of hierarchical injection channels. From these injections hierarchical and coaxial laminar flows were formed before they solidified. When the complex was flowed into the collection channel and reacted with the outer CaCl2 sheath flow in situ, a calcium alginate microfiber that had
Figure 3. a) Schematic illustration of the hierarchical capillary coaxial microfluidic device used for continuously generating hollow microfibers; b,c) digital photographs of the microfluidic device and its microstructure during the formation of the hollow microfibers; d,e) microscope image of the hollow alginate microfibers with one and two channels; f–i) cross-sectional CLSM images of the hollow microfibers with one, two, three, and four channels. All scale bars indicate 200 µm. The experimental flow rate conditions were Qcore-fluid/QNa-alginate/QCaCl2 = 1:2:15; 0.1:0.7:10; 0.6:1.8: 15; 0.4:1.8:15; 0.3:1.6: 15; 0.15:1.8:15 mL h−1, respectively in (d–i).
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Figure 4. Schematic illustrations of the multishell hollow microfibers and their corresponding microfluidic injection channels, and the cross-sectional CLSM images of the microfibers: a) with two-shell layers at the flow rate of the four fluids (Qcore-fluid/QNa-alginate-in red/QNa-alginate-in green/QCaCl2) is 0.2:0.6:1:15 mL h−1; b) with three-shell layers at the flow rate of the four fluids (Qcore-fluid/QNa-alginate-in green1/QNa-alginate-in red/QNa-alginate-in green2/QCaCl2) is 0.2:0.5:0.7:1.1:15 mL h−1; c) with Janus-compartmental shell layers; d) with four-compartmental shell layers; e) with six-compartmental shell layers. The scale bar indicates 200 µm. The specific flow rate conditions were Qcore-fluid/QNa-alginate/QCaCl2 = 0.7:0.7:15; 0.4:0.35:15; 0.5:0.4:15 mL h−1, respectively in (c–e).
the same multiple shell-layer structure as the injection flows was generated. Figure 4a and 4b show the CLSM images of two-shell hollow microfibers and three-shell hollow microfibers. It can clearly be seen that the internal architectures of these microfibers had an annular distribution of the individual compartments. The multichannel hollow microfibers and multiple-shell-layer microfibers showed physical structures similar to blood vessels in the human
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body. Thus, they might find potential applications in tissue engineering. It should be noted that different kinds of hollow microfibers with multicompartmental shell layers could also be generated using a single microfluidic device. Such a device was constructed by aligning a tapered seven-barrel injection capillary and a collection capillary coaxially inside a square capillary, as shown in Figure S8 in the Supporting Information.
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COMMUNICATION Figure 5. a) Optical images and fluorescent images of HepG2-laden microfibers during a two-week culture, the nuclei of the HepG2 cells were stained with blue fluorescent dye. b) Optical images and fluorescent images of cellular spheroids released from the microfibers corresponding to those in (a). The scale bar indicates 200 µm in (a) and 100 µm in (b). c,d) CLSM images of cell-laden microfibers with: c) Janus-compartments and d) a twoshell layer hollow structure. NIH 3T3 cells were stained with a red fluorescent dye, HepG2 cells were stained with a green fluorescent dye. The scale bar indicates 200 µm. e) Cell viability in the Janus-compartmental microfibers and the two-shell layer hollow microfibers during a one-week culture. f) Results of the cell MTT assay corresponding to (e). All data were measured at the same initial cell number and were expressed as mean ± s.d. (n ≥ 10). The specific flow rate condition was QHepG2/Q3T3/QCaCl2 = 1:1: 12 mL h−1 in (c), whereas the flow rate condition was Qcore-fluid/QHepG2/Q3T3/QCaCl2 = 0.33:1:1:12 mL h−1 in (d).
The central channel of the injection capillary was filled with the core fluid of CaCl2, while the six surrounding channels could be filled with middle fluids of Na-alginate with different actives. Figure 4c–e represents typical cross-sectional confocal images of the resultant hollow microfibers with Januscompartmental shell layers, four-compartment shell layers, and six-compartment shell layers. Because of this ability to
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control the detailed microstructure, these microfibers might find applications in the replication of complex tissues found in living organisms. To demonstrate the feasibility of using such microfibers in the application of biomedical engineering, HepG2 cells or NIH 3T3 cells were dispersed into the Na-alginate solution and the mixtures were pumped into the microfluidic devices to
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fabricate cell-laden microfibers. The optical images and SEM images (Figure S9, Supporting Information) show that the cells were uniformly and separately dispersed in the resultant microfibers, which provided a 3D microenvironment for cell growth. The viability of the encapsulated cells was near to 80%. This high biocompatibility is due to the simplicity of the encapsulation method, which depends only on aqueous–aqueous gelation instead of using oil, heat, or UV light. Thus, this method has little influence on the resulting material properties. The growth process of the HepG2 cells in the microfibers was investigated and recorded continuously for two weeks, as shown in Figure 5 and Figure S10 (Supporting Information). It was found that the HepG2 cells proliferated and gradually formed cellular spheroids in the microfibers (Figure 5a). The cellular spheroids could be released from the microfibers by using alginate lyase to selectively digest the calcium alginate (Figure 5b). These cellular spheroids are believed to be capable of reflecting the in vivo physiology of tumors more realistically than two-dimensional cell cultures, and have found important applications in drug discovery. Although many methods have been developed to form cellular spheroids, it is difficult to control their size and to produce large numbers of them. In contrast, our method could generate cell-laden fibers continuously, and thus correspondingly form large numbers of cellular spheroids. In addition, because the spheroids originated from single cells and their size increased with incubation time, the size of the cellular spheroids could be controlled by releasing them at different times. Cell co-culture experiments were also investigated in the microfibers. For this purpose, the HepG2 mixed solution and NIH 3T3 mixed solution were used to fabricate Janus-compartmental microfibers and two-shell hollow microfibers. To observe the distribution of cells in the microfibers, the two kinds of cells were stained green and red, respectively, before the encapsulation. From CLSM images of the resultant cell-laden microfibers (Figure 5c,d), it could be seen that there is a distinct boundary at the interface between the HepG2 and NIH 3T3 cells, both in the Janus-compartmental microfibers and the two-shell hollow microfibers. The cells in these structural microfibers still had a high viability before and after long-term culture (Figure 5e). However, with further research, it was found that the co-culture of HepG2 and NIH 3T3 cells in the two-shell hollow microfibers resulted in a better performance than that of the Janus-compartmental microfibers (Figure 5f). This could be ascribed to the hollow microstructure of the former, which provided not only a larger contact surface for the two cells but also a more effective route for absorption of the medium and cell metabolism. Thus, these complex structural microfibers might provide novel 3D cell co-culture models for biological research. In conclusion, we have presented a novel, the multiplelaminar-flow microfluidic method for the scalable formation of alginate microfibers with tunable, morphological, structural, and chemical features. The primary applications of these heterotypic microfibers for cell co-culture and 3D growth were also demonstrated. It is worth mentioning that when these cells were encapsulated in multiple-shell hollow microfibers, the individual cells had an annular distribution in the microfibers, and this indicates potential value of these materials for the construction of artificial blood vessels. In addition, other kinds of cell-laden microfibers could also be used as biological models
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in organisms, thus we expect that these microfibers will lead to a host of novel applications in tissue engineering.
Supporting Information Supporting Information is available from the Wiley Online Library or from the author.
Acknowledgements This research was supported by the National Natural Science Foundation of China (Grants 91227124, 21105011 and 21327902), the research Fund for the Doctoral Program of Higher Education of China (20120092130006), the Program for Changjiang Scholars and Innovative Research Team in University (IRT1222), and the Technology Invocation Team of Qinglan Project of Jiangsu Province. Y.J.Z. acknowledges the Program for New Century Excellent Talents in University. Received: February 19, 2014 Revised: April 24, 2014 Published online:
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Bioinspired Multicompartmental Microfibers from Microfluidics Yao Cheng, Fuyin Zheng, Jie Lu, Luoran Shang, Zhuoying Xie, Yuanjin Zhao,* Yongping Chen,* and Zhongze Gu* Microfibers are long, thin, and flexible materials. They are useful for the creation of various functional three-dimensional (3D) objects, including clothes and other architectures by folding, bundling, reeling, and weaving.[1–6] These microfibers have also been attractive for creating complex 3D tissues in vitro because the hierarchical structures of the human body include various types of fiber-shaped 3D cellular constructions, such as blood vessels,[7–9] muscle fibers,[10] nerve bundles,[11] hepatic cord structures,[12] and similar anatomical structures.[13–15] However, the fiber-shaped structures in plants and animals usually possess compositional and topographical properties that vary spatiotemporally on the microscale.[16] Thus, to engineer these linear tissues, preparation of heterogeneous cellular microfibers with similar microstructures is essential for regulating the direction and distribution of cell growth.[17,18] Several methods have recently been developed to fabricate microfibers continuously, such as electrospinning, wet spinning, and melt spinning.[19–25] However, most of the microfibers generated by these techniques have a homogeneous chemical composition and structure, and the creation of continuous microfibers with spatiotemporally tunable chemical compositions and structures remains a great challenge. Microfluidics has created exciting avenues of scientific and engineering research into the synthesis of functional materials.[26–30] Microfluidics is the technology that deals with the precise control and manipulation of small quantities of fluids constrained in microchannels of small cross-sectional dimensions.[31] Because of this feature, microfluidics is considered to be a promising candidate for continuous fabrication of functional microfibers. A series of research projects has demonstrated the feasibility of engineering continuous microfibers with single or multiple compositions and structures by using coaxial polydimethylsiloxane (PDMS) microfluidic
Dr. Y. Cheng,[+] Dr. F. Y. Zheng,[+] Dr. J. Lu, Dr. L. R. Shang, Prof. Z. Y. Xie, Prof. Y. J. Zhao, Prof. Z. Z. Gu State Key Laboratory of Bioelectronics Southeast University Nanjing 210096, China E-mail: [email protected]; [email protected] Prof. Y. P. Chen School of Energy and Environment Southeast University Nanjing 210096, China E-mail: [email protected] [+]These authors contributed equally to this work
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channels.[32–35] These systems have broken through several barriers associated with conventional methods.[36,37] However, due to the limitation of generating microchannels of fixed dimensions by current microfluidic techniques, it has not been possible to generate heterogeneous microfibers with complex morphologies and tunable compositions to the same extent as those present in natural organisms. Here, we present a one-step, continuous process for the scalable formation of microfibers with desired features using novel capillary microfluidics. Our method employs immediate microfiber gelation from multiflows of sodium alginate (Na-alginate) solution and calcium chloride (CaCl2) solution at the orifice of an injection capillary, which is coaxially aligned with a collection capillary inside a square capillary (Figure 1a–c). As these flows had a low Reynolds number, they mainly formed laminar flows in the microfluidic channels and only slowly diffusively mixed across the adjacent streams (Figure S1, detailed description in the Supporting Information). Thus, the resultant microfibers had the same structure as the injection flows. Homogeneous structures as well as microfibers possessing anisotropic multicompartmental body, core, and shell compositions could be generated by varying the injection capillary design. To our knowledge, such microfibers have not previously been reported. Furthermore, we have also explored the potential use of these heterotypic microfibers for tissue-engineering applications by creating multifunctional fibers with a spatially controlled encapsulation of cells. The proposed microfibers may enable the construction of fiber-shaped tissues in natural organisms. In a typical experiment, the inner phase of the Na-alginate precursor was pumped through the tapered capillary, while the CaCl2 stream was pumped in the same direction through the region between the inner capillary and the outer square capillary. Facilitated by hydrodynamic focusing, a 3D coaxial sheath flow stream around the flow of the precursor was formed at the merging of both flows. Thus, because of fast diffusioncontrolled ion cross-linking, a hydrogel microfiber was generated in situ. It is worth mentioning that by changing the flow rates, the microfibers could form four kinds of flow patterns, including clogging, wavy microfibers, straight microfibers, and spiral microfibers, in the collection channel (Figure S2, Supporting Information). Among these, the flow pattern of straight microfibers was desired (as is explained in the Supporting Information). Figure 1d and 1e show photographs of calcium alginate microfibers resulting from this. The microfibers had a cylindrical structure and high uniformity. The diameter and speed of generation of the microfibers was investigated. The speed can be increased by simply increasing the core
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Figure 1. a) Schematic illustration of the capillary coaxial microfluidic device used for continuously generating simple microfibers. b,c) Digital photographs of the microfluidic device and its microstructure during the formation of the microfibers. d) Digital image of continuous microfibers in a vessel. e) Microscope image of the alginate microfibers, the scale bar indicates 200 µm. f) Relationship between the diameter of the microfibers and the flow rates of the Na-alginate solution (blue color, with a fixed CaCl2 flow rate of 15 mL h−1) and calcium chloride solution (red color, with a fixed Na-alginate flow rate of 5 µL min−1); the orifice diameter of the tapered injection capillary of the microfluidic device is 50 µm.
and sheath flow rates. In a specific microfluidic device (with a tapered tip of 50 µm and a collection channel of 580 µm), the relationships between the microfiber diameter and the core and sheath flow rates are shown in Figure 1f. It was found that because of core-flow solidification, the practical diameters of the microfibers were larger than those theoretically calculated from the pure fluids (Figure S3, Supporting Information). The diameter of the microfibers increased in a flow-rate-dependent manner of the Na-alginate flow, showing an increase of about 90 µm as the sheath flow rate increased from 1 to 35 µL min−1. In contrast, as the CaCl2 sheath flow rate increased from 4 to 50 mL h−1, the fiber diameter decreased by 80 µm on average. A larger adjustment range of the microfiber diameter could be achieved by using different microfluidic devices (Figure S4, Supporting Information). To fabricate multicomponent microfibers, our basic microfluidic device underwent a series of transformations. As shown in Figure 2a, instead of the single-channel injection capillary previously described, a tapered two-barrel capillary was used as the injection channel and coaxially inserted into the collection capillary. Two fluids of Na-alginate solution mixed with different dyes were simultaneously pumped into the injection capillary channels using the same flow rate. Because of their low Reynolds number, they mainly formed a laminar flow and no obvious diffusive mixing occurred across the two color streams before being solidified (Figure 2b,c). When they were flowed into the collection channel and reacted with the CaCl2 sheath flow in situ, a two-compartment Janus calcium alginate microfiber formed stably, similar to the formation process of the single-compartment microfibers. Figure 2d represents typical cross-sectional confocal laser scanning microscopy (CLSM) images of the Janus microfibers. It was found that the internal architecture of the microfiber displayed an extremely sharp
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boundary at the interface between the two compartments, suggesting minimal mass transfer between the compartments. By replacing the two-barrel injection capillary with a four- or six-barrel injection capillary, four- and six-compartment calcium alginate fibers could correspondingly be generated, as shown in Figure 2e and 2f. Each compartment of the microfibers could have its own chemical composition, which is determined by the chemical composition of the initial injecting solutions. Thus, the multicompartmental microfibers have great potential applications in creating multiactive encapsulations. The volume ratio of each compartment could also be modulated by controlling the flow rates of the injection fluids. Hollow alginate microfibers were also fabricated using a microfluidics capillary with hierarchical injection channels, as indicated in Figure 3a. When three fluids, namely, a core fluid (CaCl2), a middle fluid (Na-alginate), and an outer sheath fluid (CaCl2), were introduced into the microfluidic device (Figure 3b), a hollow microfiber was generated in two stages. During the first stage, a coaxial flow of the core fluid and middle fluid was generated and the primary gelation process started along the inner wall of the hollow microfiber. In the second stage, the two coaxial fluids merged with the calcium ion sheath flow, and together they formed a three-layered coaxial flow. The gelation occurred simultaneously at both the inner and outer layers of the middle Na-alginate fluid. The three-layered coaxial flow moved through the collection capillary and the solidified alginate hollow microfiber was continuously generated and extruded (Figure 3c). The hollow microfibers achieved were of uniform shell thickness, as shown in Figure 3d and 3f. Both the diameter of the hollow microfibers and the thickness of the microfiber shell could be modulated by controlling the physical parameters of the fluids or tuning the flow rates of the fluids (as shown in Figure S5, Supporting
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COMMUNICATION Figure 2. a) Schematic illustration of the microfluidic device used for generating Janus-compartmental microfibers. b,c) Digital photographs of the microfluidic device and its microstructure during the formation of the Janus microfibers. d–f) Schematic illustrations of the multicompartmental microfibers and their corresponding microfluidic injection channels, and the cross-sectional CLSM images of the multicompartmental microfibers: d) Janus-compartments; e) four-compartments; f) six-compartments. The scale bar indicates 200 µm. The experimental flow rate conditions were QNa-alginate/QCaCl2 = 0.5:10; 0.3:10; 0.2:10 mL h−1, respectively in (d–f).
Information). Hollow microfibers with multiple core channels could also be generated by inserting several spindle capillaries in parallel to the original injection capillary for pumping of the core fluids (Figure 3g–i and Figure S6, Supporting Information). The hollow microstructures of these microfibers are confirmed by the scanning electron microscopy (SEM) images in Figure S7 in the Supporting Information.
To fabricate hollow microfibers with multiple shell layers, a microfluidic device was constructed with an increasing number of hierarchical injection channels. From these injections hierarchical and coaxial laminar flows were formed before they solidified. When the complex was flowed into the collection channel and reacted with the outer CaCl2 sheath flow in situ, a calcium alginate microfiber that had
Figure 3. a) Schematic illustration of the hierarchical capillary coaxial microfluidic device used for continuously generating hollow microfibers; b,c) digital photographs of the microfluidic device and its microstructure during the formation of the hollow microfibers; d,e) microscope image of the hollow alginate microfibers with one and two channels; f–i) cross-sectional CLSM images of the hollow microfibers with one, two, three, and four channels. All scale bars indicate 200 µm. The experimental flow rate conditions were Qcore-fluid/QNa-alginate/QCaCl2 = 1:2:15; 0.1:0.7:10; 0.6:1.8: 15; 0.4:1.8:15; 0.3:1.6: 15; 0.15:1.8:15 mL h−1, respectively in (d–i).
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Figure 4. Schematic illustrations of the multishell hollow microfibers and their corresponding microfluidic injection channels, and the cross-sectional CLSM images of the microfibers: a) with two-shell layers at the flow rate of the four fluids (Qcore-fluid/QNa-alginate-in red/QNa-alginate-in green/QCaCl2) is 0.2:0.6:1:15 mL h−1; b) with three-shell layers at the flow rate of the four fluids (Qcore-fluid/QNa-alginate-in green1/QNa-alginate-in red/QNa-alginate-in green2/QCaCl2) is 0.2:0.5:0.7:1.1:15 mL h−1; c) with Janus-compartmental shell layers; d) with four-compartmental shell layers; e) with six-compartmental shell layers. The scale bar indicates 200 µm. The specific flow rate conditions were Qcore-fluid/QNa-alginate/QCaCl2 = 0.7:0.7:15; 0.4:0.35:15; 0.5:0.4:15 mL h−1, respectively in (c–e).
the same multiple shell-layer structure as the injection flows was generated. Figure 4a and 4b show the CLSM images of two-shell hollow microfibers and three-shell hollow microfibers. It can clearly be seen that the internal architectures of these microfibers had an annular distribution of the individual compartments. The multichannel hollow microfibers and multiple-shell-layer microfibers showed physical structures similar to blood vessels in the human
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body. Thus, they might find potential applications in tissue engineering. It should be noted that different kinds of hollow microfibers with multicompartmental shell layers could also be generated using a single microfluidic device. Such a device was constructed by aligning a tapered seven-barrel injection capillary and a collection capillary coaxially inside a square capillary, as shown in Figure S8 in the Supporting Information.
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COMMUNICATION Figure 5. a) Optical images and fluorescent images of HepG2-laden microfibers during a two-week culture, the nuclei of the HepG2 cells were stained with blue fluorescent dye. b) Optical images and fluorescent images of cellular spheroids released from the microfibers corresponding to those in (a). The scale bar indicates 200 µm in (a) and 100 µm in (b). c,d) CLSM images of cell-laden microfibers with: c) Janus-compartments and d) a twoshell layer hollow structure. NIH 3T3 cells were stained with a red fluorescent dye, HepG2 cells were stained with a green fluorescent dye. The scale bar indicates 200 µm. e) Cell viability in the Janus-compartmental microfibers and the two-shell layer hollow microfibers during a one-week culture. f) Results of the cell MTT assay corresponding to (e). All data were measured at the same initial cell number and were expressed as mean ± s.d. (n ≥ 10). The specific flow rate condition was QHepG2/Q3T3/QCaCl2 = 1:1: 12 mL h−1 in (c), whereas the flow rate condition was Qcore-fluid/QHepG2/Q3T3/QCaCl2 = 0.33:1:1:12 mL h−1 in (d).
The central channel of the injection capillary was filled with the core fluid of CaCl2, while the six surrounding channels could be filled with middle fluids of Na-alginate with different actives. Figure 4c–e represents typical cross-sectional confocal images of the resultant hollow microfibers with Januscompartmental shell layers, four-compartment shell layers, and six-compartment shell layers. Because of this ability to
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control the detailed microstructure, these microfibers might find applications in the replication of complex tissues found in living organisms. To demonstrate the feasibility of using such microfibers in the application of biomedical engineering, HepG2 cells or NIH 3T3 cells were dispersed into the Na-alginate solution and the mixtures were pumped into the microfluidic devices to
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fabricate cell-laden microfibers. The optical images and SEM images (Figure S9, Supporting Information) show that the cells were uniformly and separately dispersed in the resultant microfibers, which provided a 3D microenvironment for cell growth. The viability of the encapsulated cells was near to 80%. This high biocompatibility is due to the simplicity of the encapsulation method, which depends only on aqueous–aqueous gelation instead of using oil, heat, or UV light. Thus, this method has little influence on the resulting material properties. The growth process of the HepG2 cells in the microfibers was investigated and recorded continuously for two weeks, as shown in Figure 5 and Figure S10 (Supporting Information). It was found that the HepG2 cells proliferated and gradually formed cellular spheroids in the microfibers (Figure 5a). The cellular spheroids could be released from the microfibers by using alginate lyase to selectively digest the calcium alginate (Figure 5b). These cellular spheroids are believed to be capable of reflecting the in vivo physiology of tumors more realistically than two-dimensional cell cultures, and have found important applications in drug discovery. Although many methods have been developed to form cellular spheroids, it is difficult to control their size and to produce large numbers of them. In contrast, our method could generate cell-laden fibers continuously, and thus correspondingly form large numbers of cellular spheroids. In addition, because the spheroids originated from single cells and their size increased with incubation time, the size of the cellular spheroids could be controlled by releasing them at different times. Cell co-culture experiments were also investigated in the microfibers. For this purpose, the HepG2 mixed solution and NIH 3T3 mixed solution were used to fabricate Janus-compartmental microfibers and two-shell hollow microfibers. To observe the distribution of cells in the microfibers, the two kinds of cells were stained green and red, respectively, before the encapsulation. From CLSM images of the resultant cell-laden microfibers (Figure 5c,d), it could be seen that there is a distinct boundary at the interface between the HepG2 and NIH 3T3 cells, both in the Janus-compartmental microfibers and the two-shell hollow microfibers. The cells in these structural microfibers still had a high viability before and after long-term culture (Figure 5e). However, with further research, it was found that the co-culture of HepG2 and NIH 3T3 cells in the two-shell hollow microfibers resulted in a better performance than that of the Janus-compartmental microfibers (Figure 5f). This could be ascribed to the hollow microstructure of the former, which provided not only a larger contact surface for the two cells but also a more effective route for absorption of the medium and cell metabolism. Thus, these complex structural microfibers might provide novel 3D cell co-culture models for biological research. In conclusion, we have presented a novel, the multiplelaminar-flow microfluidic method for the scalable formation of alginate microfibers with tunable, morphological, structural, and chemical features. The primary applications of these heterotypic microfibers for cell co-culture and 3D growth were also demonstrated. It is worth mentioning that when these cells were encapsulated in multiple-shell hollow microfibers, the individual cells had an annular distribution in the microfibers, and this indicates potential value of these materials for the construction of artificial blood vessels. In addition, other kinds of cell-laden microfibers could also be used as biological models
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in organisms, thus we expect that these microfibers will lead to a host of novel applications in tissue engineering.
Supporting Information Supporting Information is available from the Wiley Online Library or from the author.
Acknowledgements This research was supported by the National Natural Science Foundation of China (Grants 91227124, 21105011 and 21327902), the research Fund for the Doctoral Program of Higher Education of China (20120092130006), the Program for Changjiang Scholars and Innovative Research Team in University (IRT1222), and the Technology Invocation Team of Qinglan Project of Jiangsu Province. Y.J.Z. acknowledges the Program for New Century Excellent Talents in University. Received: February 19, 2014 Revised: April 24, 2014 Published online:
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