Spherical Phospholipid Polymer Hydrogels for Cell Encapsulation

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Spherical Phospholipid Polymer Hydrogels for Cell Encapsulation Prepared with a Flow-Focusing Microfluidic Channel Device Tatsuo Aikawa,†,§ Tomohiro Konno,‡ Madoka Takai,‡ and Kazuhiko Ishihara*,†,‡,§ †

Department of Materials Engineering, School of Engineering, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-8656, Japan ‡ Department of Bioengineering, School of Engineering, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-8656, Japan § Center for Medical System Innovation (CMSI), The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-8656, Japan ABSTRACT: To prepare spherical polymer hydrogels, we used a flow-focusing microfluidic channel device for mixing aqueous solutions of two water-soluble polymers. Continuous encapsulation of cells in the hydrogels was also examined. The polymers were bioinspired 2methacryloyloxyethyl phosphorylcholine polymer bearing phenyl boronic acid groups (PMBV) and poly(vinyl alcohol) (PVA), which spontaneously form a hydrogel in aqueous medium via specific molecular complexation upon mixing, even when they were in cell culture medium. The microfluidic device was prepared with polydimethylsiloxan, and the surface of the channel was treated with fluoroalkyl compound to prevent sticking of the polymers on the surface. The microfluidic channel process could control the diameter of the spherical hydrogels in the range of 30−90 μm and generated highly monodispersed diameter spherical hydrogels. We found that the polymer distribution in the hydrogel was influenced by the PVA concentration and that the hydrogel could be dissociated by the addition of D-sorbitol to the suspension. The single cells could be encapsulated and remain viable in the hydrogels. The localized distribution of polymers in the hydrogel may provide an environment for modulating cell function. It is concluded that the spontaneous hydrogel formation between PMBV and PVA in the flow-focusing microfluidic channel device is applicable for continuous preparation of a spherical hydrogel-encapsulating living cell.

1. INTRODUCTION The encapsulation of living cells into a hydrophilic polymer hydrogel plays an important role in bioengineering fields such as tissue engineering,1 cell-based transplantation therapy,2,3 and single-cell analysis.4 Several methods for cell encapsulation have been reported.2,4−12 These methods can be generally classified into two groups: micronozzle dropping methods or microfluidic channel methods. The dropping method produces relatively large-sized capsules containing hundreds of cells. This method allows us to prepare a cluster of pancreas cells or islets of Langerhans cells in the hydrogel capsules and has been applied to cell-based transplantation therapy.3 By contrast, the microfluidic channel method can precisely generate microsized hydrogel capsules, and the number of encapsulated cells can be regulated.5,6,12 When two immiscible fluids such as a pair of oil and prepolymer aqueous solutions are injected into a channel with a cross-shaped junction, uniform sized hydrogel capsules are produced by the shear force generated between the two fluids.13 Regulating the flow rates of fluids allows us to control the size of the hydrogel from tens to several hundreds of micrometers in the diameter. The diameter of the hydrogel capsule defines some important properties relating to the environment around the cells such as permeability of the solute and intercellular communication. Taking advantage of these © 2011 American Chemical Society

benefits, we employed a microfluidic channel for cell encapsulation in this work. Some properties of the polymer hydrogels are also important in terms of tuning cell behavior. Various types of polymers have been employed for cell encapsulation, including naturally derived polymers,14,15 synthetic polymers,7,8 and modified natural polymers.9,10 When encapsulating living cells, polymers must be able to form a cross-link under mild conditions to avoid damaging cells. In terms of this reason, a naturally derived polymer is suitable since they can form hydrogels under relatively mild conditions via electrostatic interactions or intermolecular interaction. However, one drawback of such polymers is that it is difficult to modulate properties affecting cell behaviors. Porosity, elasticity, or types of functional groups of hydrogels relate to transportation of solute, cell migration, cell adhesion, or inducing chemical signal on the cellular membrane.16 Therefore these relationships dominate cell viability, proliferation, and differentiation. By contrast, the use Special Issue: Bioinspired Assemblies and Interfaces Received: September 26, 2011 Revised: December 12, 2011 Published: December 16, 2011 2145

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Figure 1. The chemical structure of PMBV (a), and the gelation mechanism between PMBV and PVA (b). an oil bath. The resultant product was purified by reprecipitation with a mixture of ether and chloroform (70:30 by volume) followed by dialysis against water for 7 days (Spectra/Por 7 membrane MWCO 3500; Spectrum Laboratories Inc., Rancho Dominguez, CA, USA). The yield of PMBV was 86% after the purification. The composition of the purified polymer was determined by 1H NMR (JNM-AL300; JEOL, Tokyo, Japan). The averaged molecular weight was determined by gel permeation chromatography (JASCO, Tokyo, Japan) with an OHpak SB-804 HQ column (Shodex; Showa Denko, Tokyo, Japan) using methanol/distilled water (70:30 by volume) containing 10 mM lithium bromide as the eluent. The column was calibrated with poly(ethylene glycol) standards. The chemical structure of PMBV and the gelation mechanism with PVA are shown in Figure 1. 2.2. Methods. 2.2.1. Fabrication of the Microfluidic Device. 2.2.1.1. Microfluidic Channel Mold. The epoxy resin mold for fabricating the PDMS microfluidic device was prepared by UV lithography as follows. A glass slide (S1112; Matsunami Glass, Tokyo, Japan) was washed with ethanol in a sonicator bath for 10 min and then treated with oxygen plasma at 300 W for 10 min under 100 Pa of oxygen atmosphere in a plasma generator (Plasma Reactor PR 500; Yamato Scientific, Tokyo, Japan). A defined quantity of photoreactive epoxy resin polymer (SU-8 50; Microchem, MA, U.S.A.) was coated onto the pretreated slide glass by spin coating at 500 rpm for 5 s and then at 1000 rpm for 30 s. The slide glass was heated for 10 min at 65 °C followed by 30 min at 95 °C. A specially designed photomask was attached to the slide glass, which was then irradiated with UV light at 300 W/cm2 for 30 s. After irradiation, the slide glass was heated at 95 °C for 10 min and then rinsed in an SU-8 developer (Microchem) for 10 min to dissolve the unreacted epoxy resin polymer. Finally, the SU8 mold was obtained. 2.2.1.2. PDMS Microfluidic Device. PDMS prepolymer (Silpot 184; Dow-Toray, Tokyo, Japan) and a curing agent were mixed together at a ratio of 10:1 by weight. The mixture was poured into a tray containing the fabricated SU-8 mold to a depth of ca. 5 mm and degassed under reduced pressure. The mixture was heated at 65 °C for 4 h to solidify the PDMS, and the solidified PDMS was then carefully peeled from the mold for obtaining PDMS replica. The PDMS replica was bound to a flat PDMS plate as a bottom plate immediately after oxygen plasma treatment at 85 W for 30 s under 20 Pa of oxygen atmosphere and placed in an oven at 85 °C for 2 h. The PDMS microchannel was then fluorine modified. This process is necessary to prevent adhesion of the polymer to the microchannel during preparation of the spherical hydrogels. Modification with fluoroalkyl compound was carried out under trichloro(1H,1H,2H,2Hperfluorodecyl)silane (TCFS; TCI) vapor. The PDMS microchannel was placed in a desiccator with TCFS for 18 h under 14 kPa. To confirm the modification, X-ray photoelectron spectroscopy (XPS; Kratos/Shimadzu, Kyoto, Japan) was performed using Mg Kα sources with a photoelectron takeoff angle of 90°.

of synthetic polymers allows control over the hydrogel properties through modification of the constituent polymers. In this work, we used a hydrogel formed from the bioinspired phospholipid polymer, poly(2-methacryloyloxyethyl phosphorylcholine (MPC)-co-n-butyl methacrylate (BMA)-co-4-vinylphenyl boronic acid (VPBA)) (PMBV), and poly(vinyl alcohol) (PVA) as materials for cell encapsulation (Figure 1a). The PMBV/PVA hydrogel is highly biocompatible and can be prepared under mild conditions without irradiation or heat.17,18 Moreover, the hydrogel forms spontaneously via specific molecular complexation when aqueous solutions of PMBV and PVA are mixed. The cross-linking of PMBV and PVA, which results from spontaneous reaction between a single boronic acid moiety of PMBV and two hydroxyl groups of PVA, proceeds even in cell culture medium. Importantly, the PMBV/PVA hydrogel can be dissociated by the addition of a monosaccharide, such as D-sorbitol, which has a higher binding constant to the boronic acid moiety than does PVA (Figure 1b).18−21 Here, we used a flow-focusing microfluidic channel device prepared with polydimethylsiloxan (PDMS) by soft lithography technique. Monodispersed microsized droplets of the polymers were formed by injection of oil and aqueous solutions of PMBV and PVA into the microfluidic device. Human promyelocytic leukemia (HL-60) cells were then encapsulated in the hydrogel by dispersing the cells in the polymer solution prior to injection into the microfluidic device. Controlling the size and size distribution of the spherical hydrogels and the mixing state of the polymers in the hydrogels were examined. Such cellencapsulating spherical hydrogel will be useful not only for cellbased transplantation therapy2,3 but also as a tool in cell biology research, especially for techniques involving single-cell analysis.4

2. MATERIALS AND METHODS 2.1. Materials. PMBV was synthesized by conventional radical polymerization according to the following process:17 MPC (120 mmol, synthesized as reported by us elsewhere22), BMA (40 mmol, Kanto Chemicals, Tokyo, Japan), VPBA (40 mmol, Tokyo Chemical Industry (TCI), Tokyo, Japan), and 2, 2′-azobisisobutyronitrile (2 mmol, Kanto Chemicals) were dissolved in 200 mL of ethanol (Kanto Chemicals) and poured into a 300 mL four-neck round-bottom flask equipped with a thermometer, reflux condenser, argon gas inlet, septum cap, and magnetic stirring bar. Argon gas was bubbled into the monomer solution for 20 min to deoxygenate the solution prior to polymerization. Copolymerization was performed at 60 °C for 12 h in 2146

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Figure 2. Photograph of the PDMS microfluidic channel equipped with a stainless holder (a), the structures of the microchannel (b), and the orifice (c). 2.2.2. Continuous Preparation of Spherical Hydrogels with the Microfluidic Device. PMBV and PVA (mean polymerization degree = 1000, saponification degree = 96.0 mol %, Wako Pure Chemical, Osaka, Japan) were dissolved in cell culture medium (RPMI-1640; Cat No. R8758, Sigma-Aldrich) and injected separately into two inlets of the microfluidic device (Figure 2b). Paraffin oil (Wako) containing 2.0 wt % of sorbitan monooleate (Span 80; TCI) as a surfactant was injected into the other two inlets. The injections were performed with gastight syringes (1001TLL; Hamilton Company, GR, Switzerland) using a microsyringe pump (BS-MD1001; Bioanalytical Systems, IN, U.S.A.) to control the flow rate precisely. 2.2.3. Evaluation of Polymer Distribution in the Spherical Hydrogels. The distribution of polymers in the spherical hydrogels was determined by confocal laser scanning microscopy using an LSM 510 microscope (Carl Zeiss, Oberkochen, Germany) equipped with a C-Apochromat 40×/1.2W objective lens and using a HeNe Laser (λ = 543 nm) for excitation. For these experiments, hydrogels were synthesized by replacing PMBV with its rhodamine derivative (poly(MPC-co-n-BMA-co-VPBA-co-methacryloxyethyl thiocarbamoyl rhodamine B; PMBV-R), which has a fluorescent emission wavelength of ∼550 nm. The composition of the rhodamine derivative in the PMBV-R was about 0.03 unit mole fraction. 2.2.4. Evaluation of the Dissociation Behavior of the Spherical Hydrogels. The spherical hydrogels ejected from the microfluidic device were collected into the concave of a glass slide. Oil was removed by addition of a few drops of n-hexane, and then phosphatebuffered saline containing 1.0 M D-sorbitol was added to the suspension to dissociate the spherical hydrogel. The dissociation behavior of the hydrogels was observed with an optical microscope (IX71; Olympus, Tokyo, Japan), with the dissociation time being defined as the point at which the hydrogel outline disappeared. 2.2.5. Cell Encapsulation in the Spherical Hydrogels. To demonstrate cell encapsulation in the microsized PMBV/PVA hydrogel, HL-60 cells were suspended in a PMBV/RPMI-1640 aqueous solution supplemented with 50 U/mL penicillin and 50 μg/ mL streptomycin (Invitrogen, NY, U.S.A.). Before encapsulation, 1.0 mL of the cell suspension was incubated in Live/Dead solution containing 2 μM calcein-AM and 4 μM ethidium homodimer-1 (Live/ Dead viability/cytotoxicity kit; Invitrogen) to determine the viability of encapsulated cells by fluorescence microscopy.

The mole fraction of the MPC units in the polymer was less than 0.50, and the polymer had poor solubility in water.23,24 To secure sufficient water-solubility for preparing the hydrogel, we set the mole fraction of MPC units above 0.50 in the polymer. To maintain the hydrogel structure homogeneously, the distribution of the cross-linking points is important. When the composition of VPBA units in the polymer was more than 0.20, the gelation rate was too high to obtain a homogeneous gel structure. Thus, we determined the composition of the each monomer unit in the PMBV. When PMBV and PVA solutions were separately injected into the PDMS microfluidic device, the solutions mixed at a junction in the microfluidic device, and the flux was pinched off into droplets by shear force derived from the stream of oil (Figure 3a). The droplets gelated immediately after mixing at the junction.

3. RESULTS AND DISCUSSION 3.1. Continuous Preparation of Spherical Hydrogels with the Microfluidic Device. We obtain a water-soluble PMBV as a random copolymer with the following mole fractions: MPC, BMA, and VPBA were 0.71, 0.13, and 0.16, respectively (Figure 1a). The composition of PMBV was decided in terms of the following: controllability of polymerization, solubility in water, and homogeneity of resulting hydrogel. The BMA conld regulate the overall polymerization rate during polymerization. When the polymerization was carried out without the BMA, the polymerization rate of MPC and VPBA was extremely high. As a result, the molecular weight and composition of each monomer unit were hardly controlled. The composition of the MPC units in polymer was optimized from the viewpoint of the solubility of the polymer in water.

Figure 3. Optical microscopic images of polymer solution flow in the PDMS microfluidic channel device with fluorine modification (a), and without fluorine modification (b). Scale bar = 200 μm.

Preliminary experiments revealed that droplet formation was inhibited by adhesion of the polymer to the surface of the microchannel. The accumulation of polymer forms a bump on the channel surface, and droplets that collide with the bump are disrupted (Figure 3b). Fluorine modification of the PDMS microchannel was performed to prevent the adhesion of 2147

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the hydrogels was dependent on the flow-rate ratio (Qc/Qd), where Qc and Qd are the flow rate of the oil and the polymer solution, respectively. The diameter of the hydrogels decreased with increasing Qc/Qd. When Qc/Qd was less than 1.0, the flux of the polymer solution was elongated at the junction, which allowed the formation of fiber-shaped hydrogels as well as spherical ones. The coefficient of variation (Cv) of the diameter of the spherical hydrogel was less than 5.0% at all Qc/Qd (where Cv is the standard distribution divided by the mean diameter). Thus, the use of the microfluidic device in the preparation of spherical hydrogels allows us to control the size precisely and to produce highly uniform spherical hydrogels. 3.2. Evaluation of Polymer Distribution in the Spherical Hydrogel. The concentration of PMBV-R in the spherical hydrogels was evaluated by confocal fluorescence microscopy (Figure 5). The PMBV-R was homogeneously distributed in the hydrogels formed with 5.0 wt % PMBV-R and 1.0 wt % PVA (Figure 5a) but not in hydrogels with other polymer compositions (Figure 5b−f). In the case of the hydrogels with 5.0 wt % PMBV-R and 1.0 wt % PVA, the viscosity increased slowly during the gelation reaction. Accordingly, PMBV-R and PVA could mix by diffusion of molecules within the droplet. By contrast, the gelation reaction was more rapid in hydrogels with higher PVA concentrations, which caused viscosity to be drastically increased and prevented adequate mixing of PMBV-R and PVA during the diffusion process. When droplets migrate through a microchannel with low Reynolds number, mixing of fluids within droplets is induced by diffusion and vortex.25 If a winding channel can be installed in our device, it would enhance mixing of fluids within droplets.26 The state of mixing in the spherical hydrogels may influence the dissociation behavior, mechanical property, or the diffusion of molecules in the hydrogel and thus may also have a physiological effect on cells encapsulated in the hydrogel. 3.3. Evaluation of Dissociation Behavior of the Spherical Hydrogels. All of the spherical PMBV/PVA hydrogels could be dissociated within 10 min of addition of D-sorbitol solution, regardless of their PMBV and PVA composition. By contrast, approximately 1 h incubation was

polymer and subsequent disruption of droplets. To confirm the existence of fluoroalkyl compound on the surface of PDMS after the modification process, we performed XPS measurement. Atomic ratios of fluorine to silicon (F/Si) before and after the surface modification were 0.6 and 21.4, respectively. This result indicated that fluoroalkyl compound was bound on the PDMS surface. As expected, surface modification with the fluoroalkyl compound prevented adhesion of PMBV/PVA hydrogels to the microchannel surface and allowed the formation of droplets (Figure 3a). Another important issue for the preparation of PMBV/PVA spherical hydrogels is the need to ensure that the polymers remain separate before droplets are pinched off. Even if PMBV and PVA come in contact for a few seconds, the polymers gelate rapidly, the viscosity of the flux increases drastically, and the gelated flux cannot be pinched off by shear force. To shorten the contact time between PMBV and PVA prior to pinching off, the flow rate of each polymer solution should be greater than 0.1 μL/min. Figure 4 shows the relationship between the flow-rate ratio and the diameter of the spherical hydrogels. The diameter of

Figure 4. The relationship between flow-rate ratio (Qc/Qd) and the diameter of the spherical PMBV/PVA hydrogels, where Qc and Qd are the flow rate of the oil and the polymer solution, respectively. Qc was set at 5.0 μL/min (open circle) or 10.0 μL/min (filled circle). The error bar indicates standard distribution.

Figure 5. Confocal microscopic images of spherical PMBV-R/PVA hydrogels. The area enclosed by the dashed line indicates the outline of the hydrogel. Bright area in the spherical hydrogels indicates the region distributing PMBV-R. [PMBV-R]/[PVA] (wt %/wt %) = 5.0/1.0 (a), 5.0/2.5 (b), 5.0/5.0 (c), 10.0/1.0 (d), 10.0/2.5 (e), and 10.0/5.0 (f). Scale bar = 50 μm. 2148

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days.2,17,27 In addition, the PMBV/PVA hydrogels can be implanted in vivo in tissue surrounding tendons.28 Thus, the PMBV/PVA hydrogels is suitable for encapsulation of cells. The proliferation of the HL-60 cells depended on the cell density. We could not observe significant proliferation of the HL-60 cells due to low cell density on the culture plate, after dissociation of the spherical hydrogel to collect the HL-60 cells and plating them on the culture plate. However, Live/Dead assay clearly indicated that the HL-60 cells were viable after the collection. We are currently evaluating the functions of the cells encapsulated in the PMBV/PVA hydrogels by examining their gene expression. The results of these studies will be reported in the near future.

required to dissociate a bulky PMBV/PVA hydrogel with the same composition and ca. 1 mL volume. The shorter dissociation time is likely due to the spherical hydrogels having a larger specific surface area than the bulky hydrogel and consequently having more cross-linked sites exposed to Dsorbitol. The dissociation time was lengthened by increasing concentrations of PVA, which may be explained by more complex entanglement of polymers and larger amount of hydrogen bonding between hydroxyl groups of PVA in the hydrogels containing higher amounts of PVA. 3.4. Cell Encapsulation in the Spherical Hydrogels. To control the number of cells in the each hydrogel, we altered several parameters, such as the density of cells in the PMBV and the Qd and Qc values. When the density of cells was 1.0 × 107 cells/mL in the PMBV portion and Qd was 1.0 μL/min, the distribution of encapsulating single cells in one spherical hydrogel depended on the diameter, which was changed by Qc. That is, the distribution of encapsulating single cells in one spherical hydrogel was 31% and 26% at 40 μm and 52 μm diameter of the spherical hydrogel, respectively. Although empty hydrogels were observed more frequently in the case of 40 μm diameter than in the case of of 52 μm, the smaller hydrogel was proper to encapsulate single cells (Figure 6a). On

4. CONCLUSIONS We have demonstrated the preparation of a spherical cytocompatible hydrogel by molecular assembly based on the mixing of PMBV and PVA solutions in a flow-focusing microchannel device. The modification of the microchannel surface with fluoroalkyl compound was necessary for continuous successful preparation of the spherical hydrogels. Regulating the flow-rate ratio between the oil phase and the polymer solution phase could readily control the size of the spherical hydrogels, and the size distribution of the spherical hydrogels generated was narrow. The spherical PMBV/PVA hydrogel could be dissociated within 10 min in monosaccharide aqueous solutions. This property will be useful for recovering cells from the spherical hydrogels under mild conditions. Changing the PVA concentration was shown to regulate the polymer distribution in the hydrogels, which is expected to facilitate fine-tuning of the mechanical and permeation properties of the hydrogels. Finally, we encapsulated living single cells within the spherical hydrogels. This result suggests that the PMBV/PVA spherical hydrogels prove to be useful for cell therapy or for applications such as single-cell analysis.

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AUTHOR INFORMATION

Corresponding Author

*Mailing address: Department of Materials Engineering, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 1138656, Japan. Tel.: +81 3 5841 7124; fax: +81 3 5841 8647. Email address: [email protected].

Figure 6. Fluorescence microscopic images of spherical PMBV/PVA hydrogels encapsulating HL-60 cells. The hydrogels with two different mean diameters, 40 μm (a) or 52 μm (b), were prepared by changing Qc between 30.0 and 40.0 μL/min, but Qd was kept constant at 1.0 μL/min. During preparation, cells were dispersed in the PMBV aqueous portion at concentrations of 1 × 107 cells/mL. Viable HL-60 cells emitted green fluorescence by staining with calcein-AM. Scale bar = 100 μm.

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ACKNOWLEDGMENTS A part of this work was supported by a Grant-in-Aid for Scientific Research on Innovative Areas ‘‘Nanomedicine Molecular Science’’ (No. 2306) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. T.A. was supported by the Center for Medical System Innovation (CMSI) program, The University of Tokyo, during conducting this research.

the other hand, the larger hydrogels induced multicell encapsulation (Figure 6b). When the diameter of the hydrogel was smaller than 40 μm, the frequency of empty hydrogels was rather increased. To control of number of cells in a spherical hydrogel strictly, we should further examine the design of microfluidic channel and the dispersion state of cells. The viability of encapsulated cells was 96%, demonstrating that neither the PMBV/PVA hydrogel itself nor the process of preparing the spherical hydrogel had an adverse effect on the cells. Indeed, as previously reported, the PMBV/PVA hydrogels possess excellent long-term cytocompatibility, with more than 90% of the cells remaining viable after immobilization for 8

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REFERENCES

(1) Brandl, F.; Sommer, F.; Goepferich, A. Rational design of hydrogels for tissue engineering: Impact of physical factors on cell behavior. Biomaterials 2007, 28, 134−146. (2) Boninsegna, S.; Toso, R. D.; Monte, R. D. Alginate microspheres loaded with animal cells and coated by siliceous layer. J. Sol-Gel Sci. Technol. 2003, 26, 1151−1157. (3) Rabanel, J. M.; Banquy, X.; Zouaoui, H.; Mokhtar, M. Progress technology in microencapsulation method for cell therapy. Biotechnol. Prog. 2009, 25, 946−961.

2149

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(4) Novak, R.; Zeng, Y.; Shuga, J.; Venugopalan, G.; Fletcher, D. A.; Smith, M. T.; Mathies, R. A. Single-cell multiplex gene detection and sequencing with microfluidically generated agarose emulsion. Angew. Chem., Int. Ed. Engl. 2011, 123, 410−415. (5) Tan, W.; Takeuchi, S. Monodisperse alginate hydrogel microbeads for cell encapsulation. Adv. Mater. 2007, 19, 2696−2701. (6) Morimoto, Y.; Tan, W.; Tsuda, Y.; Takeuchi, S. Monodisperse semi-permeable microcapsules for continuous observation of cells. Lab Chip 2009, 9, 2217−2223. (7) Koh, W.; Revzin, A.; Pishko, M. V. Poly(ethylene glycol) hydrogel microstructures encapsulating living cells. Langmuir 2002, 18, 2459−2462. (8) Panda, P.; Ali, S.; Lo, E.; Chung, G. B.; Hatton, A. T.; Khademhosseini, A.; Doyle, P. S. Stop-flow lithography to generate cell-laden microgel particles. Lab Chip 2008, 8, 1056−1061. (9) Reza, A. T.; Nicoll, S. B. Characterization of novel photocrosslinked carboxymethylcellulose hydrogels for encapsulation of nucleus pulposus cells. Acta Biomater. 2010, 6, 179−186. (10) Sakai, S.; Ito, S.; Ogushi, Y.; Hashinoto, I.; Hosoda, N.; Sawae, Y. Enzymatically fabricated and degradable microcapsules for production of multicellular spheroids with well-defined diameters of less than 150 μm. Biomaterials 2009, 30, 5937−5942. (11) Nicodemus, G. D.; Bryant, S. J. Cell encapsulation in biodegradable hydrogels for tissue engineering applications. Tissue Eng., Part B: Rev. 2008, 14, 149−165. (12) Konry, T.; Dominguez-Villar, M.; Baecher-Allan, C.; Hafler, D. A.; Yarmush, M. L. Droplet-based microfluidic platforms for single T cell secretion analysis of IL-10 cytokine. Biosens. Bioelectron. 2011, 26, 2707−2710. (13) Zhao, C.; Middelberg, A. P. J. Two-phase microfluidic flows. Chem. Eng. Sci. 2011, 66, 1394−1411. (14) Rowey, J. A.; Madlambayan, G.; Mooney, D. J. Alginate hydrogels as synthetic extracellular matrix materials. Biomaterials 1999, 20, 45−53. (15) Ouyang, W.; Chen, H.; Jones, M. L.; Metz, T.; Haque, T.; Martoni, C.; Prakash, S. Artificial cell microcapsule for oral delivery of live bacterial cells for therapy: Design, preparation, and in-vitro characterization. J. Pharm. Pharm. Sci. 2004, 7, 315−324. (16) Saul, J. M.; Williams, D. F. Hydrogel in regenerative medicine. In Principles of Regenerative Medicine; Atala, A., Lanza, R., Thomson, J. A., Nerem, R., Eds.; Elsevier: London, 2011; pp 637−661. (17) Konno, T.; Ishihara, K. Temporal and spatially controllable cell encapsulation using a water-soluble phospholipid polymer with phenylboronic acid moiety. Biomaterials 2007, 28, 1770−1777. (18) Xu, Y.; Jang, K.; Konno, T.; Ishihara, K.; Mawatari, K.; Kitamori, T. The biological performance of cell-containing phospholipid polymer hydrogels in bulk and microscale form. Biomaterials 2010, 31, 8839−8846. (19) Fujita, N.; Shinkai, S.; James, T. D. Boronic acids in molecular self-assembly. Chem. Asian J. 2008, 3, 1076−1091. (20) Springsteen, G.; Wang, B. A detailed examination of boronic acid−diol complexation. Tetrahedron 2002, 58, 5291−5300. (21) Yan, J.; Springsteen, G.; Deeter, S.; Wang, B. The relationship among pKa, pH, and binding constants in the interactions between boronic acids and diolsIt is not as simple as it appears. Tetrahedron 2004, 60, 11205−11209. (22) Ishihara, K.; Ueda, T.; Nakabayashi, N. Preparation of phospholipid polymers and properties as hydrogel membranes. Polym. J. 1990, 22, 355−360. (23) Ishihara, K.; Iwasaki, Y.; Nakabayashi, N. Polymeric lipid nanosphere consisting of water-soluble poly(2-methacryloyloxyethyl phosphorylcholine-co-n-butyl methacrylate). Polym. J. 1999, 31, 1231− 1236. (24) Konno, T.; Watanabe, J.; Ishihara, K. Enhanced solubility of paclitaxel using water-suluble and biocompatible 2-methacryloyloxyethyl phosphorylcholine polymers. J. Biomed. Mater. Res., Part A 2003, 65, 209−214.

(25) Bringer, M. R.; Gerdts, C. J.; Song, H.; Tice, J. D.; Ismagilov, R. F. Microfluidic system for chemical kinetics that rely on chaotic mixing in droplets. Philos. Trans. R. Soc. London, A 2004, 362, 1087−1104. (26) Muradoglu, M.; Stone, H. A. Mixing in a drop moving through a serpentine channel: A computational study. Phys. Fluids 2005, 17, 073305. (27) Xu, Y.; Sato, K.; Mawatari, K.; Konno, T.; Jang, K.; Ishihara, K.; Kitamori, T. A microfluidic hydrogel capable of cell preservation without perfusion culture under cell-based assay condition. Adv. Mater. 2010, 22, 3017−3021. (28) Ishiyama, N.; Moro, T.; Ohe, T.; Miura, T.; Ishihara, K.; Konno, T.; Ohyama, T.; Kimura, M.; Kyomoto, M.; Saito, T.; Nakamura, K.; Kawaguchi, H. Reduction of peritendinous adhesions by hydrogel containing biocompatible phospholipid polymer MPC for tendon repair. J. Bone Jt. Surg. Am. 2011, 93A (2), 142−149.

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