Technical Note pubs.acs.org/ac
Three-Dimensional Cell Culture and Drug Testing in a Microfluidic Sidewall-Attached Droplet Array Shi-Ping Zhao,† Yan Ma,† Qi Lou,† Hong Zhu,‡ Bo Yang,‡ and Qun Fang*,† †
Institute of Microanalytical Systems, Department of Chemistry and Innovation Center for Cell Signaling Network, Zhejiang University, Hangzhou, 310058, China ‡ College of Pharmaceutical Sciences, Zhejiang University, Hangzhou, 310058, China S Supporting Information *
ABSTRACT: Three-dimensional (3D) cell culture provides an effective way over conventional two-dimensional (2D) monolayer culture to more closely imitate the complex cellular organization, heterogeneity, and interactions as well as tissue microenvironments in vivo. Here we present a novel dropletbased 3D cell culture method by using droplet array attached on the sidewall of a PDMS piece. Such an arrangement not only avoids cells from adhering on the chip surface for achieving 3D cell culture in nanoliter-scale droplets, but also facilitates performing multiple operations to cells in droplets, including cell suspension droplet generation, drug treatment, and cell staining with a capillary-based liquid handling system, as well as in situ observation and direct scanning with a confocal laser scanning microscope. We optimized the system by studying the effects of various conditions to cell culture including droplet volume, cell density and fabrication methods of the PDMS pieces. We have applied this system in the 3D culture of HepG2 cells and the stimulation testing of an anticancer drug, doxorubicin, to 3D cell spheroids.
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lead to limitations in applying to assay or screening with scarce or expensive samples or reagents. The emergence of microfluidic chip technique has provided a miniaturized platform for 3D cell culture with low consumption, high cell spheroid formation efficiency, integrated devices,15 and better control to spheroid sizes16,17 and flows in spatial and temporal domains, which could mimic in-vivo-like microenvironments with high precision and throughput.16 So far, a variety of different microfluidic methods18−20 have been developed to perform 3D cell culture for forming cell spheroids in microfluidic systems, which could be divided into three types, hanging-drop,21,22 microstructure array,16,23−26 and droplet-based microfluidics.27,28 As shown in Figure 1a, the microfluidic hanging-drop devices have a similar structure as conventional systems in which cells aggregate to the concave bottom of a drop hung on the bottom of a substrate by gravity, and tend to form cell spheroids by the natural organization through cell−cell attachment and production of extracellular matrix (ECM). However, the structure of hanging drop systems with drops hung on the substrate bottom obstructs the subsequent assay operations to these drops, such as addition of drugs or labeling dyes, as well as real-time observation of indrop cells. Although some access holes could be fabricated on the substrate to facilitate subsequent operation, 21 the
ell culture under three-dimensional (3D) mode can better mimic the actual complex environment in vivo than conventional two-dimensional (2D) culture mode, in terms of extracellular matrix components, cell-to-cell and cell-to-matrix interaction.1−3 Compared with 2D culture mode, the results of drug screening under 3D mode are closer to clinical tests and can provide more realistic prediction for safety and risk assessment. Researchers have developed a variety of 3D cell culture methods to simulate the in vivo characteristics of physiological tissues. Among these methods, there are two types of 3D culture modes: cell spheroid-based mode in which cells spontaneously self-aggregate and gradually form spheroids,4 and gel-based mode under which cells are cultured in hydrogel scaffolds, such as extracellular matrix, agarose and so on.5−7 The cell spheroid-based culture is one of the widely used 3D culture methods, which has advantages of simple device structure and ease in operation. In addition, a tumor cell spheroid formed under 3D mode can naturally and more closely mimic an avascular solid tumor in generating substance gradient along the spheroid radius such as nutrients, metabolites, and oxygen, as well as stimulated drugs. Over the past decades, several methods for cell spheroid formation have been developed, including hanging drop method,8−10 nonadhesive culture surface,11 microfabricated structure,12 spinner flask cultures,13 and rotary bioreactors.14 These methods have been widely applied in current biological and medical researches. However, the reagent consumptions of these methods are usually in the microliter range, which may © 2017 American Chemical Society
Received: June 12, 2017 Accepted: September 8, 2017 Published: September 8, 2017 10153
DOI: 10.1021/acs.analchem.7b02267 Anal. Chem. 2017, 89, 10153−10157
Technical Note
Analytical Chemistry
Figure 1. Different microfluidic 3D cell culture methods for cell spheroid formation: (a) Hanging drop; (b) Continuous flow-based droplet system; (c) Microstructure array; (d) Droplet array attached on the sidewall of a PDMS substrate.
Figure 2. (a) Illustration of cell culture and drug testing process in the present sidewall-attached droplet array: (1) Cell seeding; (2) Formation of cell spheroid in a droplet; (3) Addition of the drug; (4) Addition of fluorescence dye to stain the cell spheroid. (b) Photograph of a sidewall-attached droplet array with red and green dye droplets. (c) Detail view of an array of red dye droplets attached on the sidewall of a PDMS piece. (d) Schematic diagram of a sidewallattached droplet array system with cell spheroids forming in droplets.
consumptions of these systems are still in the range of tens microliters. The droplet-based microfluidic systems27,28 (Figure 1b) can generate gel droplets with high throughput for forming cell spheroids in droplets. However, these systems need to couple with other microchip systems to carry out the subsequent operation of cell assay. For the microstructure array method16,23−26 (Figure 1c), U-shaped or cubic-shaped microstructures are usually fabricated in a microchip to trap cells into the pocket of each microstructure for 3D culture through the actions of fluid flow and gravity. Since continuous perfusion is generally needed in this type of systems, the reagent consumptions are usually in the microliter or even milliliter range.23,24 Here we present a novel droplet-based 3D cell culture mode using nanoliter-scale droplet array attached on the sidewall of a PDMS substrate (Figure 1d), which allows the flexible and convenient implementation of a series of 3D cell assay operations including droplet generation, drug treatment, cell staining, in situ observation, and direct scanning with a confocal laser scanning microscope. For such a novel in-droplet 3D cell culture mode, we investigated the effects of various conditions to cell culture including droplet volume, cell density, and fabrication methods of the PDMS pieces. We applied this system in the 3D culture of HepG2 cells and the stimulation testing of anticancer drug doxorubicin to 3D cell spheroids.
Procedure. In-Droplet 3D Cell Culture. Before cell experiment, the device was autoclaved for sterilization, and the PDMS pieces were exposed to UV light for 12 h to achieve the bonding with the Petri dish surface. To avoid droplet evaporation and offer a gas-permeable microenvironment for in-droplet cell culture, the PDMS pieces were covered with 5 mL Fluorinert oil (FC-40, 3M, St. Paul, MN).29 Droplet array of cell suspension were formed on the sidewall of the PDMS pieces using the droplet handling mode of sequential operation droplet array (SODA).29−35 Briefly, a definite volume of HepG2 cell suspension with a cell density of 2 × 106 cells/mL was first aspirated into the tapered capillary probe with hydrophobic inner and outer surfaces under control of the syringe pump, and deposited on the sidewall of the PDMS pieces under a microscope to form a droplet (Figure 2a1). Multiple droplets were sequentially generated as the above procedure on both sidewalls of each PDMS pieces to form a droplet array, and the distance between adjacent droplets was about 3 mm. After a 3-day culture, tumor cells of HepG2 formed cell spheroids in the nanoliter-scale droplets attached on the sidewall of the PDMS pieces (Figure 2a2). Anticancer Drug Assay. In this work, doxorubicin (Dox, Toronto Research Chemicals Inc., Toronto, Canada) was used as the anticancer drug in the assay. A series of solutions of doxorubicin with concentrations of 2 × 10−4, 2 × 10−5, 2 × 10−6, 2 × 10−7, 2 × 10−8, 2 × 10−9, and 2 × 10−10 M were prepared with a 0.1 M doxorubicin stock solution. The culture medium without doxorubicin was used as control. After a 3-day culture, HepG2 tumor cells formed cell spheroids in the droplets (Figure 2a2). A 500 nL doxorubicin solution was added to each tested droplet to stimulate the cell spheroids for 1 day (Figure 2a3). After the drug stimulation, a 200 nL PBS solution with two 2% (v/v) fluorescence dyes, calcein-AM and EthD-1 from a live/dead cell kit (Life technologies, Carlsbad, CA), was added to each droplet (Figure 2a4). The droplet array was incubated at 37 °C for 1 h, and then the tumor cell spheroids in droplets were scanned with a spinning disk
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EXPERIMENTAL SECTION Setup of the Droplet Array Device. The microfluidic droplet array device is composed of multiple polydimethylsiloxane (PDMS) pieces located in a Petri dish and a capillary probe (530 μm i.d., 690 μm o.d., Refine Chromatography Co., Yongnian, China) with a tapered tip (tip size: ∼120 μm i.d. and ∼150 μm o.d.) connected with a syringe pump (PHD 2000, Harvard Apparatus, Holliston, MA). For the fabrication of PDMS pieces, ∼6.5 g PDMS prepolymer (Sylgard 184, Dow Corning, Midland, MI) was prepared at 10:1 (w/w) ratio, degassed and added into a Petri dish (6 cm diameter) for curing at 80 °C for 2 h. The solidified PDMS with a thickness of ∼4 mm was cut into pieces by a scalpel with a piece width of 0.5−1 cm. The PDMS pieces were placed in a Petri dish with a distance of 3−10 mm between adjacent pieces. Figure 2 shows the schematic diagram and images of the device. 10154
DOI: 10.1021/acs.analchem.7b02267 Anal. Chem. 2017, 89, 10153−10157
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Analytical Chemistry
composite force of the droplet’s gravity and adhesion forces can balance its upward buoyancy. When the droplet volume was larger than 3 μL, the droplets could not be stably attached to the sidewall of the PDMS pieces, exhibiting a floating tendency to the top surface of the PDMS piece, since the adhesive forces of these larger droplets to the PDMS surface could not resist against their buoyancy. For droplets with volumes in the range of 1 to 3 μL, although attached droplets could be formed on the PDMS sidewall, they easily move to the top surface of the PDMS piece during the subsequent cell assay operation process also due to the insufficient adhesive force on the PDMS sidewall. With droplet volumes in the range of 500 to 100 nL, the droplets could stably attached on the PDMS piece surface without evidently moving and splitting in the experiment process, even with the shocks produced by multiple falling of the whole device from 1 cm height. Thus, such a phenomenon of droplet adhering on PDMS sidewall has evident scaling effect, especially for nanoliter-scale rather than microliter-scale droplets. However, in the subsequent in-droplet cell culture experiment, for 100 or 300 nL droplets, an obvious droplet evaporation phenomenon was observed during a 3-day incubation. For this situation, a liquid supplement operation to these droplets is needed during the culture process to ensure the success rate of forming cell spheroids in droplets. To simplify the experimental operation, we chose 500 nL as the volume of droplets. Cell Density. We tested the influence of different cell densities of 5 × 105, 1 × 106, 2 × 106, 3 × 106, and 4 × 106 cells/mL. Some typical results are shown in Figure S1. The cell density of 2 × 106 cells/mL demonstrated both sufficient cell viability of 87% and success rate of 66% (144 droplets from three independent experiments) and, thus, was adopted in the followed experiments. Fabrication Methods of the PDMS Pieces. In this system, the droplet array was designed to be attached on the sidewall of the PDMS pieces. We chose PDMS material due to its good biocompatibility and ease in fabrication.29 We tested the effects of the casting and cutting fabrication methods for the PDMS pieces, especially for their sidewall, on the performance of droplet attachment and 3D cell culture. The PDMS pieces fabricated by the casting method using a glass cuvette as a mold had smooth sidewall surface, resulting in the difficulty in observing cell spheroids in droplets with a microscope due to the light reflection (Figure S2). The PDMS pieces produced by the cutting method with a scalpel had a relatively rough sidewall surface, which not only were capable of carrying droplets to perform 3D cell culture, but also avoided the optical interference of the sidewall to cell observation. 3D Spheroid Culture. With the optimized conditions, namely, 500 nL droplets, 2 × 106 cells/mL cell density, and a 3day incubation without changing culture medium, 3D cell culture could be performed in the present droplet array system with an occurrence rate for cell spheroids in droplets of 66%. In the experiment, cell spheroids of liver cancer cells, HepG2, we successfully formed. Figure 3a illustrates the formation process of a spheroid of HepG2 tumor cells in a droplet during 4-day culture period, and Figure 3b shows the phase contrast and confocal microscope images of HepG2 cells spheroids formed in different droplets. We also carried out an experiment using longer cell culture time of 5 days without medium replacement, and the result is shown in Figure S3. Although the cell viability showed a slight
confocal microscope (IX81, Olympus, Tokyo, Japan). In the fluorescence images, calcein AM is shown in green (Ex/Em: 488 nm/525 nm), and EthD-1 is shown in red (Ex/Em: 561 nm/600 nm). The z-stack images obtained by the confocal microscope were used to calculate cell viability with the software Bitplane Imaris (Andor Technology, Belfast, Northern Ireland).
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RESULTS AND DISCUSSION System Design. Hanging-drop8−10,21,22 is a widely used technique for 3D cell culture capable of forming cell spheroids in drops. It is based on the use of microliter-scale drops hung on a substrate and performs cell culture at the bottom of each drop without any risk of cell adhesion to the substrate. However, such an arrangement results in the inconvenience in performing subsequent operations to drops, such as addition of drugs and dyes, as well as real-time observation of in-drop cells. Although some access holes could be fabricated on the substrate to facilitate subsequent operation,10,21 the consumptions of these systems are still in the range of tens of microliters. In this work, we used a different droplet “hanging” mode by attaching droplets on the substrate sidewall instead of its bottom as in conventional8−10 and other microfluidic21,22 methods, utilizing the evident interface adhesion effect of substrate sidewall to nanoliter-scale droplets. This droplet attaching mode could also avoid cells from adhering on the substrate surface as the previous hanging drop methods8−10,21,22 and, thus, achieve 3D cell culture in droplets. Furthermore, such a special arrangement opens both the spaces above and under the droplets, which could significantly facilitate the handling operation to droplets with a capillary probe. By using the sequential operation droplet array (SODA)29−35 droplet manipulation strategy, multistep operations, including droplet generation and reagent additions, required for cell assay were conveniently achieved with the present system. To avoid the evident evaporation of nanoliterscale droplets during cell culture and assay process, we used a gas-permeable perfluorocarbon oil (Fluorinert FC-40 from 3M, St. Paul, MN) to cover the droplets, which has been demonstrated to have good biocompatibility with cell culture.29 This measure did not interfere with the droplet handling operation due to the semiopen property of the oil-covered droplets which allows the capillary probe to handle droplets through the oil. With the present method, a series of nanoliterscale droplets could be generated on substrates to form a droplet array for performing large scale and high-throughput screening. System Optimization. For achieving 3D cell culture with the formation of cell spheroids in sidewall-attached droplets, we studied the influences of various conditions on the cell culture performance including droplet volume, cell density, and fabrication methods of the PDMS pieces. Droplet Volume. In the present system, we used the scaling phenomenon that microdroplets can be adhered on a solid surface to build the droplet array. We first studied the stability of various droplets with different volumes of 10 μL, 3 μL, 1 μL, 500 nL, 300 nL, and 100 nL on the sidewall of a PDMS piece. An attached aqueous droplet immersed in oil is subjected to three forces, gravity, buoyancy of the oil, and adhesion force of the PDMS surface to the droplet. Due to the higher density (1.8 g/mL) of the FC-40 oil than that of water (1.0 g/mL), an essential condition to obtain stable attached droplets is that the 10155
DOI: 10.1021/acs.analchem.7b02267 Anal. Chem. 2017, 89, 10153−10157
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Figure 3. (a) Microscopic images showing the formation process of HepG2 cell spheroid in a sidewall-attached droplet during a 4-day culture period; (b) Microscopic images of HepG2 cell spheroids in 10 droplets attached on the sidewall of a PDMS piece after a 3-day culture. Scale bars: 100 μm.
Figure 4. Cell spheroid-based anticancer drug testing. (a) Typical microscopic images of tumor cell spheroids stimulated by different concentrations of doxorubicin. The green cells are live cells stained by calcein AM and the red ones are dead cells stained by EthD-1. The experiment for each drug concentration was repeated three times using six cell droplets per assay. (b) Comparison the results of doxorubicin testing using 2D and 3D cell culture modes. The data shown are the average results of three independent experiments. Statistical significance is determined by two-tailed Student’s t-test; *p < 0.01, **p < 0.001. Scale bars: 100 μm.
reduction with the time, the cell viability of HepG2 spheroids in droplets still remained higher than 80%, which is sufficient long for performing cell spheroid-based drug testing. Cell Spheroid-Based Anticancer Drug Testing. Benefiting from the flexible multistep operation ability of the present system, we further applied the 3D cell droplet array system to the testing of an anticancer drug, doxorubicin. As shown in Figure 2a, HepG2 cells formed cell spheroids in nanoliter-scale droplets attached on the sidewall of the PDMS chip after a 3day culture. Solutions of doxorubicin which inhibits RNA and DNA synthesis with serial concentrations were administrated to the droplets, and then the droplets were incubated at 37 °C and 5% CO2 for 1 day, followed by cell staining and confocal laser microscope scanning. The confocal microscope z-stack images of cell spheroids exposed to doxorubicin with serial concentrations were used to determine the cell viability. Figure S4 shows the confocal images of a typical cell spheroid in a sidewall-attached droplet. The step length of the z-stack images was 6 μm, by which the diameter of the cell spheroid could be deduced as ca. 130 μm. The dead cells stained with EthD-1 (red) mostly locate on the surface rather than the core of the cell spheroid, indicating that the outside cells of the cell spheroid were stimulated by the drug more strongly than the core cells. Figure 4a shows a series of typical live/dead staining images of Dox-treated spheroids obtained with different doxorubicin concentrations. The repeated experiment results (Figure 4b) shows that doxorubicin exhibits evident dosedependent decreases in cell viability. We also carried out a comparison experiment using 96-well plates as 2D cell culture mode, the results show that the cell viabilities of 3D spheroids are higher than those of 2D cultured cells when the doxorubicin concentration is higher than 100 nM (Figures 4b and S5). This indicates that HepG2 cells are more resistant to doxorubicin under the 3D culture mode than 2D mode, which can be attributed to the barrier function of the outer cells in the
spheroid to the diffusion of the drug toward the inside core cells. Such a result is consistent with the previous literature36 and demonstrates the advantage of 3D culture in mimicking in vivo conditions more closely than 2D mode.
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CONCLUSION In summary, the present system provides a novel droplet-based 3D cell culture mode which enables flexible and convenient multiple operations for 3D cell culture and assay, such as droplet generation, drug treatment, cell staining, and cell observation, owing to its more open operation space. By coupling with an automated liquid handling robot, the present method could be applied in massive and high throughput drug screening. In addition, this droplet array system is simple to build, without the use of microfabricated chips, making it easy to be applied in routine biological laboratories. In addition to the drug testing as demonstrated in this work, the present system could also provide an effective platform for cell biological research, and medical diagnosis, due to its properties of simple system setup, low consumption and cost, and flexible operation.
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.7b02267. 10156
DOI: 10.1021/acs.analchem.7b02267 Anal. Chem. 2017, 89, 10153−10157
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Analytical Chemistry
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(22) Frey, O.; Misun, P. M.; Fluri, D. A.; Hengstler, J. G.; Hierlemann, A. Nat. Commun. 2014, 5, 4250. (23) Fu, C.-Y.; Tseng, S.-Y.; Yang, S.-M.; Hsu, L.; Liu, C.-H.; Chang, H.-Y. Biofabrication 2014, 6, 015009. (24) Ziolkowska, K.; Stelmachowska, A.; Kwapiszewski, R.; Chudy, M.; Dybko, A.; Brzozka, Z. Biosens. Bioelectron. 2013, 40, 68−74. (25) Kwapiszewska, K.; Michalczuk, A.; Rybka, M.; Kwapiszewski, R.; Brzozka, Z. Lab Chip 2014, 14, 2096−2104. (26) Liu, W.; Yan, M.; Zhao, L.; Ma, C.; Li, T.; Xu, J.; Wang, J. Lab Chip 2016, 16, 4106−4120. (27) Chan, H.-F.; Zhang, Y.; Ho, Y.-P.; Chiu, Y.-L.; Jung, Y.; Leong, K.-W. Sci. Rep. 2013, 3, 3462. (28) Agarwal, P.; Zhao, S.; Bielecki, P.; Rao, W.; Choi, J. K.; Zhao, Y.; Yu, J.; Zhang, W.; He, X. Lab Chip 2013, 13, 4525−4533. (29) Du, G.-S.; Pan, J.-Z.; Zhao, S.-P.; Zhu, Y.; den Toonder, J. M. J.; Fang, Q. Anal. Chem. 2013, 85, 6740−6747. (30) Zhu, Y.; Zhang, Y.-X.; Cai, L.-F.; Fang, Q. Anal. Chem. 2013, 85, 6723−6731. (31) Jin, D.-Q.; Zhu, Y.; Fang, Q. Anal. Chem. 2014, 86, 10796− 10803. (32) Zhu, Y.; Zhu, L.-N.; Guo, R.; Cui, H.-J.; Ye, S.; Fang, Q. Sci. Rep. 2014, 4, 5046. (33) Zhu, Y.; Zhang, Y.-X.; Liu, W.-W.; Ma, Y.; Fang, Q.; Yao, B. Sci. Rep. 2015, 5, 9551. (34) Ma, Y.; Pan, J.-Z.; Zhao, S.-P.; Lou, Q.; Zhu, Y.; Fang, Q. Lab Chip 2016, 16, 4658−4665. (35) Liu, W.-W.; Zhu, Y.; Feng, Y.-M.; Fang, J.; Fang, Q. Anal. Chem. 2017, 89, 822−829. (36) Yip, D.; Cho, C. H. Biochem. Biophys. Res. Commun. 2013, 433, 327−332.
Cell culture; Anticancer Drug Assay; Influence of cell density on 3D cell cuture; Figures S1−S5 (PDF).
AUTHOR INFORMATION
Corresponding Author
*Phone: +86-571-88206771. E-mail:
[email protected]. ORCID
Qun Fang: 0000-0002-6250-252X Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We greatly appreciate the financial support of the Natural Science Foundation of China (Grants 21435004 and 21227007), and the Major National Science and Technology Programs (Grant 2013ZX09507005).
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REFERENCES
(1) Asthana, A.; Kisaalita, W. S. Drug Discovery Today 2012, 17, 810− 817. (2) Pampaloni, F.; Reynaud, E. G.; Stelzer, E. H. K. Nat. Rev. Mol. Cell Biol. 2007, 8, 839−845. (3) Yamada, K. M.; Cukierman, E. Cell 2007, 130, 601−610. (4) Fennema, E.; Rivron, N.; Rouwkema, J.; van Blitterswijk, C.; de Boer, J. Trends Biotechnol. 2013, 31, 108−115. (5) Shin, Y.; Han, S.; Jeon, J. S.; Yamamoto, K.; Zervantonakis, I. K.; Sudo, R.; Kamm, R. D.; Chung, S. Nat. Protoc. 2012, 7, 1247−1259. (6) Lee, G. H.; Lee, J. S.; Wang, X.; Hoon Lee, S. Adv. Healthcare Mater. 2016, 5, 56−74. (7) Marasso, S. L.; Puliafito, A.; Mombello, D.; Benetto, S.; Primo, L.; Bussolino, F.; Pirri, C. F.; Cocuzza, M. Microfluid. Nanofluid. 2017, 21, 29. (8) Timmins, N. E.; Dietmair, S.; Nielsen, L. K. Angiogenesis 2004, 7, 97−103. (9) Kelm, J. M.; Timmins, N. E.; Brown, C. J.; Fussenegger, M.; Nielsen, L. K. Biotechnol. Bioeng. 2003, 83, 173−180. (10) Tung, Y.-C.; Hsiao, A. Y.; Allen, S. G.; Torisawa, Y.-S.; Ho, M.; Takayama, S. Analyst 2011, 136, 473−478. (11) Friedrich, J.; Seidel, C.; Ebner, R.; Kunz-Schughart, L. A. Nat. Protoc. 2009, 4, 309−324. (12) Napolitano, A. P.; Chai, P.; Dean, D. M.; Morgan, J. R. Tissue Eng. 2007, 13, 2087−2094. (13) Xu, F.; Xu, L.; Wang, Q.; Ye, Z.; Zhou, Y.; Tan, W.-S. BioMed Res. Int. 2014, 2014, 1−10. (14) Lei, X.-H.; Ning, L.-N.; Cao, Y.-J.; Liu, S.; Zhang, S.-B.; Qiu, Z.F.; Hu, H.-M.; Zhang, H.-S.; Liu, S.; Duan, E.-K. PLoS One 2011, 6, e26603. (15) Mehta, G.; Hsiao, A. Y.; Ingram, M.; Luker, G. D.; Takayama, S. J. Controlled Release 2012, 164, 192−204. (16) Patra, B.; Peng, C.-C.; Liao, W.-H.; Lee, C.-H.; Tung, Y.-C. Sci. Rep. 2016, 6, 21061. (17) Patra, B.; Chen, Y.-H.; Peng, C.-C.; Lin, S.-C.; Lee, C.-H.; Tung, Y.-C. Biomicrofluidics 2013, 7, 054114. (18) Sung, K. E.; Beebe, D. J. Adv. Drug Delivery Rev. 2014, 79−80, 68−78. (19) Alessandri, K.; Sarangi, B. R.; Gurchenkov, V. V.; Sinha, B.; Kiessling, T. R.; Fetler, L.; Rico, F.; Scheuring, S.; Lamaze, C.; Simon, A.; Geraldo, S.; Vignjevic, D.; Domejean, H.; Rolland, L.; Funfak, A.; Bibette, J.; Bremond, N.; Nassoy, P. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 14843−14848. (20) Kuo, C.-T.; Chiang, C.-L.; Chang, C.-H.; Liu, H.-K.; Huang, G.S.; Huang, R. Y.-J.; Lee, H.; Huang, C.-S.; Wo, A. M. Biomaterials 2014, 35, 1562−1571. (21) Oliveira, M. B.; Neto, A. I.; Correia, C. R.; Rial-Hermida, M. I.; Alvarez-Lorenzo, C.; Mano, J. F. ACS Appl. Mater. Interfaces 2014, 6, 9488−9495. 10157
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