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Micropillar arrays enabling single microbial cell encapsulation in hydrogels† Kyun Joo Park,a Kyoung G. Lee,*b Seunghwan Seok,a Bong Gill Choi,bc Moon-Keun Lee,b Tae Jung Park,bd Jung Youn Park,e Do Hyun Kim*a and Seok Jae Lee*b Single microbial cell encapsulation in hydrogels is an important task to find valuable biological resources for human welfare. The conventional microfluidic designs are mainly targeted only for highly dispersed spherical bioparticles. Advanced structures should be taken into consideration for handling such aggregated and non-spherical microorganisms. Here, to address the challenge, we propose a new type of cylindrical-shaped micropillar array in a microfluidic device for enhancing the dispersion of cell clusters and the isolation of individual cells into individual micro-hydrogels for potential practical
Received 16th January 2014, Accepted 18th February 2014 DOI: 10.1039/c4lc00070f
applications. The incorporated micropillars act as a sieve for the breaking of Escherichia coli (E. coli) clusters into single cells in a polymer mixture. Furthermore, the combination of hydrodynamic forces and a flow-focusing technique will improve the probability of encapsulation of a single cell into each hydrogel with a broad range of cell concentrations. This proposed strategy and device would be a useful
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platform for genetically modified microorganisms for practical applications.
Introduction Cell-based technology has been employed to find valuable bio-entities from a variety of cellular species, such as animal and human cells, bacteria, yeast, insect, and plant cells, for research and industry.1–3 For practical purposes, it is essential to obtain the target cells from their heterogeneous cell population.4 As a result, cell isolation and encapsulation of single cells in 3D hydrogels have become a significant task to utilize their capability to produce biopharmaceuticals, including enzymes, antibodies, proteins, peptides, DNA, and metabolites for human welfare.5,6 So far, a great improvement in the a
Department of Chemical & Biomolecular Engineering, KAIST, 291 Daehak-ro, Yuseong-gu, Daejeon 305-701, Republic of Korea. E-mail: [email protected] b Center for Nanobio Integration & Convergence Engineering (NICE), National Nanofab Center, 291 Daehak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea. E-mail: [email protected], [email protected] c Department of Chemical Engineering, Kangwon National University, Samcheok 245-711, Republic of Korea d Department of Chemistry, Chung-Ang University, Seoul 156-756, Republic of Korea e National Fisheries Research and Development Institute, 216 Haean-ro, Gijang-up, Gijang-gun, Busan 619-705, Korea † Electronic supplementary information (ESI) available: Optical, fluorescent image of microparticles, detailed dimensions of chip designs, microscopic images of cell dispersion in LB medium, depleted polymer effect on cell aggregation, images of cell distribution in device, droplet size distribution histogram, cell captured droplet generation image, break up forces value with respect to cell aggregation size, details on the force calculation, and cell concentrations are presented. See DOI: 10.1039/c4lc00070f
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handling and isolation of single cells has been made using a combination of microfluidic systems, hydrogels, and fluorescence-activated cell sorting systems (FACSs).7,8 Those methods opened a new door to overcome traditional technical limitations, such as the requirement for external filters, and other bench-top equipment in complicated processes for high-throughput single-cell analysis and sorting.9 In microfluidic systems, techniques including T-junction droplet generation,9,10 droplet trapping arrays,5,10 flow-focusing droplet generation,11–14 direct cell printing,15 and droplet break up using microgroove structures9 have been developed. These techniques have also demonstrated their capability to isolate and encapsulate the cells for subsequent applications. Although the majority of invaluable biomaterials and biopharmaceuticals are discovered from non-spherical microbial cells, these approaches are mostly optimized for spherical cells and low cell concentration. Moreover, heterogeneous gene expression and rapid cellular growth rate lead to inevitable high cell concentrations and these factors combine to hamper the currently available high-throughput sorting systems and the utilization of their potential contribution in the discovery of useful bio-entities.16 In the material aspect, polymer-based hydrogels are the most popular materials to mimic an in vivo environment for cell culturing and provide hydrophilic, and non-toxic characteristics.12,17 It also possesses numerous pores for the easy access of media to maintain cell viability and further cultivation.18 However, polymers openly disturb the surface
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charges of cells and lead to serious cellular aggregation and sedimentation issues.19,20 Moreover, this inevitable phenomenon decreases the single level cell-encapsulation efficiency as well as producing non-uniform sizes of hydrogel particles. In consequence, a new breakthrough is required to adopt the benefits from both microfluidic systems and hydrogels for the handling of non-spherical biological entities, such as rod-shaped bacterial cells and concave disc-shaped red blood cells.21 Recently, it was reported that geometrical microstructures (i.e. pillar structures) in microfluidic devices can induce significant flow deformation and migration of microparticles or cells.22,23 Based on the addressed challenges and adopting structural benefits from previous reports, the incorporation of microstructures into a microfluidic device could be an alternative approach to separate,24,25 isolate,21 and trap26 the agglomerated microbials. Additionally, the currently developed droplet-based microfluidic system provides an ideal platform for isolation of cells in hydrogels. Herein, we develop an advanced concept of encapsulating non-spherical microbial single cells into hydrogel particles using the combination of micropillar arrays and a flow-focusing technique, by the continuous breaking up of microbial cell-clusters. For further demonstration, E. coli is used as a model of a non-spherical cell due to the ease with which its genes can be modified, a fast growth rate, and the availability of a complete library of network interactions between protein complexes.27 The different dimensions of the pillar arrays correspond to the different sizes of cell clusters, which result from the presence of polymers. The pillar arrays allow us to isolate cells from the clusters and increase the single-cell encapsulation efficiency. Of particular interest, we propose that the external force will enable the continuous break up of cell clusters, resulting in the production of more than 70% of single cells into individual hydrogel particles. By using the as-prepared microfluidic platform, we were able to demonstrate the applicability of the device for a wide range of cell concentrations for the isolation of cells and producing single-cell encapsulated hydrogels for potential practical applications.
Materials and methods Materials The SU-8 photoresist and developer solution were obtained from Microchem. The poly(ethylene glycol) diacrylate (PEGDA), potassium persulfate (KPS), D-sorbitol, N,N,N′,N′tetramethylethylenediamine (TEMED), and ethanol were purchased from Sigma-Aldrich. The polydimethylsiloxane (PDMS) (Dow Corning), Abil EM 90 (Degussa), and Grapeseed oil (G-oil, Beksul) were used as received. Microfluidic device fabrication Microfluidic devices in this research were fabricated using conventional soft lithography and a PDMS molding technique. Once the photoresist structure was successfully prepared by a typical photolithography process, it was used as the master mold for PDMS casting. The PDMS mixture was
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poured into the master mold and cured at 70 °C for 2 h. The PDMS layer was peeled off from the mold and the inlet and outlet were punched out for further connection. After the curing process, the PDMS-based microfluidic device was carefully cleaned using ethanol and deionized (DI) water to remove remaining fragments or dust from the surface of the device. The structured side of PDMS was bonded with a flat PDMS layer using a corona-discharge treatment and placed in the oven at 70 °C for 1 h. Cell preparation E. coli BL21(DE3) [F− ompT hsdSB (rB− mB−) gal dcm (DE3)] was used as a host strain, expressing an enhanced green fluorescent protein (EGFP). Recombinant E. coli cells harboring pET–6His–EGFP from a previous report17 were cultivated in a 250 mL flask containing Luria-Bertani (LB) medium supplemented with ampicillin of 50 μg mL−1 in a shaking incubator (200 rpm) at 37 °C. At an OD600 of 0.4, isopropyl-D-thiogalactopyranoside (IPTG) was added to a final concentration of 1 mM to induce the gene expression. Then, cells were further cultivated for 6 h and harvested by centrifugation. Cell growth was monitored by measuring the absorbance at 600 nm with a UV/Vis spectrophotometer (Optizen POP Nano Bio). Microdroplet generation A uniform size and shape of microdroplet was achieved using a flow-focusing method.17 The microfluidic device and other components including a tygon tube (0.2 in I.D., 0.6 in O.D.), syringe pumps, and a syringe were used in the experiment. The dispersed phase was a composition of different concentrations of E. coli in LB medium (0.5 mL), PEGDA (0.3 g), KPS (12.5 mg), D-sorbitol (4 mg), and DI water (0.5 mL). The PEGDA concentration for cell encapsulation, 23 wt%, adopted was the same as previous E. coli encapsulated hydrogel research.17 Additionally, the continuous phase was a mixture of G-oil and 2 wt% of Abil EM 90. Both the dispersed phase and the continuous phase were injected into inlets of the microfluidic device. In this study, Abil EM 90 was used as a surfactant to stabilize the microdroplets and prevent the fabricated droplet merging. The microdroplets were generated in the dripping regime and had a monodisperse size. The diameter size of the microdroplets was around 45 μm, under 3 μL min−1 of dispersed phase flow rate and 4 μL min−1 of continuous phase flow rate conditions. The generation of microdroplets was observed by a charge-coupled device camera (Eclipse Ti-S, Nikon, Japan) and carefully investigated using a high speed camera (Phantom V7.3). Polymerization of microdroplets The polymerization slowly started as soon as monomers were mixed with KPS, which is the initiator for chemical crosslinking. The generated droplets were collected in the mixture of G-oil and TEMED. In this work, TEMED chemically accelerated the free-radical crosslinking of the PEGDA. Once the polymerization was completed, the synthesized particles were centrifuged at 3 000 rpm for 20 s, followed by washing with isopropyl
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alcohol (IPA) and DI water to remove the excess amounts of Goil and other residues. The isolated particles were suspended in LB medium for further investigation (Fig. S1†).
the inlet. The final obtained microdroplets were collected in TEMED which accelerated the polymerization of PEGDA for further conversion into a hydrogel.
Results and discussion
Micropillar arrays in the channel
Design of overall experimental system
The microfluidic device is composed of three distinctive cylindrical-shaped micropillar zones and a flow-focusing region for monodisperse droplet generation. It has been reported that the cylindrical-shaped pillars disturb the flow pattern around pillars.22,23 Moreover, each geometrical dimension of the pillar arrays was selected based on the different sizes of the E. coil aggregations in the device (Table S1†). The detailed design and dimensions of the device and their corresponding scanning electron microscopy (SEM) images are illustrated in Fig. 2 and S2.† The first micropillar zone had pillar diameters of 100 μm, with a pillar gap distance of 50 μm. Compared to the first zone, the diameter of pillars in the second and third micropillar zones were 50 μm, but more densely packed, with narrow gaps of 25 and 10 μm, respectively. After passing through the entire micropillar zone which had a 600 μm width, the channel width is dramatically decreased down to 180 μm. The monodisperse microdroplets were simply generated by passing through the 50 μm orifice in the device. For an easier understanding of the micropillar array structures in the device, they were investigated using SEM. The obtained images, and the shape and dimensions of the pillars, showed that the PDMS microstructures were successfully replicated from the master mold as intended and properly incorporated in the device.
The different features of the cylindrical-shaped micropillarembedded microfluidic device were adapted to enhance the dispersion for non-spherical cells. The device and microstructures were fabricated using soft-lithography and PDMS. The device (Fig. 1a) is composed of two distinct functional regions for cell dispersion and encapsulation as follows: (1) micropillar zones for the breaking up of cell clusters (Fig. 1b) and (2) a flow-focusing droplet manipulator for encapsulating the single cells (Fig. 1c). By combining the two regions in a microfluidic device, it could consecutively break up cell-clusters into single cells and each cell could be encapsulated in microdroplets. For the hydrogel fabrication, PEGDA, as monomer, with a high concentration of the E. coli mixture and oil with surfactant were separately injected into the microfluidic device, as shown in Fig. 1. The flow rates of the PEGDA with cell mixture and oil were separately controlled by syringe pumps. The microdroplets were generated at the flow-focusing region after the PEGDA and cell mixture solution passed through three different stages of micropillar zones which were closely installed to
Polymer induced cell aggregation In the previous studies, cell aggregation may not have been critically discussed or observed because of no polymers being present in the medium and low inter-cellular interaction
Fig. 1 Schematic diagrams of the experimental system and hypothesis. (a) The reagents are injected into the microfluidic device by syringe pumps. The hypothesis schematics show the effect of (b) the micropillar arrays to split cell-clusters and (c) the encapsulation of single cells into the microdroplets using the flow-focusing technique.
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Fig. 2 A schematic illustration of the micropillar arrays in the microfluidic device and their corresponding SEM images. All scale bars are 100 μm.
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forces due to low cell concentration (105–107 cells mL−1).28,29 Recently, the combination of cell and polymer induced cell agglomeration has become critical and has needed to be solved for practical applications.30 To investigate the effect of polymers (i.e. PEGDA) on E. coli cells, the optical microscopy images of cells were taken under different polymeric concentrations and analyzed as shown in Fig. 3a–f. The small black dots in optical microscopic images are indicated as E. coli in LB media. Without the presence of polymers, most of the E. coli was properly dispersed, and no aggregation was observed, as shown in Fig. 3a and S3a.† By slowly increasing the amount of PEGDA up to 13 wt%, bacteria cells started to flocculate and form relatively small cell clusters in the media (Fig. 3b and S3b†). Furthermore, larger cell cluster formation was more clearly observed in the media under 23 wt% of PEGDA (Fig. 3c and S3c†). To understand and observe the polymer induced aggregation effects more clearly, the selected area from each image (red square boxes) was carefully processed and analyzed using image processing software, Image J. Each color and brightness difference in the 3D roughness from Fig. 3d–f implies the influence of polymers on the degree of cellular agglomeration. The bright blue color and smooth surface from the image represent highly dispersed cells in the media as shown in Fig. 3d. After introducing polymers, the surface roughness of the color profile images gradually increased and changed into dark blue first (Fig. 3e) then to purple (Fig. 3f). These transitions are mainly accounted for by the high level of cellular agglomeration in the media. The heterogeneous size of the cell clusters, in the range of 2 and 153 μm (Table S1†), disrupted the generation of monodisperse
Fig. 3 Microscopic images of cells in LB media (a) without PEGDA, (b) with 13 wt% of PEGDA, and (c) with 23 wt% of PEGDA. (d–f) Degree of aggregation analysis was done by measuring 3D roughness, which correspond to the red square boxes in each image through image processing. The moment of cell-cluster encapsulation in the microdroplets (g) before and (h) after passing though the orifice using no micropillar incorporated microfluidic device. Scale bars in (a–c) are 100 μm.
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microdroplets in the flow focusing region. Therefore, the combination of microorganism and polymers would inevitably generate non-uniform sizes of microdroplets (Fig. 3g and h), as well as hampering single-cell encapsulation for additional analysis and applications. From the observation and previous reports, the severe flocculation of cells is possibly explained in two aspects as follows: (1) the partially attached polymers on the cell surface disturb the net charge balance of the cell surface,19,20 and/or (2) the PEG-induced depletion attraction.31,32 The disruption of the net charge balance occurs as the introduced polymers in the media are affected by the net balance between the van der Waals (vdW) attraction and electrostatic repulsion, which can be understood from the Derjaguin, Landau, Verwey and Overbeek (DLVO) theory.31,33 Another possibility is the PEG-based polymers are depleted and the sudden decrease of their concentration brings an osmotic force and leads to the cellular agglomeration as illustrated in Fig. S4.†34,35 Overall, the combination of net charge disruption and depletion forces induce random cells to form a cluster, to become a seed for the sequential attachment of other co-existing cells in the same environment to minimize the energy.36
Cell cluster break up Distinct microstructures were employed for the effective dispersion of the bacteria clusters in the media. It demonstrates a sequential cellular break up through the continuous collision between micropillars and E. coli as shown in Fig. 4. As soon as the clusters were injected into the first micropillar zone, they collided with pillars and split into smaller size clusters (Fig. 4b–d). In this zone, the initial cell clusters were much bigger compared to the pillar gaps (50 μm) and separated from each other by the flow stream in the channel. These continuous collisions were repeated in the second and third micropillar zones and reduced the cluster size into single cells (Fig. 4e–f). These sequential processes for enhancing the cell dispersion can be understood from the combination of cluster collision, drag force, and elongation force. The flocculated cells resulted from relatively weak cell to cell interactions.35,37 As previously discussed, the vdW and depletion forces between cells are critically dependent on the distance of each cell. In this case, the closest neighbor cells show relatively strong depletion forces while the vdW and depletion forces between inner cells and outer cells get weaker as the size of cluster increases. This phenomenon led to continuous breaking cell clusters and consequently, it is more easily separated into single cells through providing such external forces. The repeatable breaking up phenomenon can be understood by the collapse of the force balance between the adhesion force and break up force. A detailed mechanism of this phenomenon is schematically demonstrated in Fig. 5. The observed cell break up was classified as two types of mechanism. Firstly, the drag and collision forces from the disperse phase flow were mainly accounted for by the continuous impact of
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then local laminar shear stress takes over the continuous break up. Both of these mechanisms are based on the flowing of the clusters around the micropillar arrays. Details on the force calculations are discussed and given in the ESI.†
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Micropillar arrays for single-cell encapsulation
Fig. 4 (a) Picture of the microfluidic device and (b–d) optical microscopic images of the first microstructural zone under different time lines. Photographs of further cell dispersion at (e) the second and (f) third microstructural zones. All the scale bars are 100 μm.
clusters with the micropillars along the flow direction (Fig. 5a–b). Secondly, the hydrodynamic disrupting force was another major factor for further breakup (Fig. 5c–d). The hydrodynamic force led to sequential elongation repeatedly,
Fig. 5 Schematic illustration of two break up mechanisms of (a) before and (b) after collision and elongation induced (c) before and (d) after break up.
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To investigate the influence of the cylindrical pillars on the cell dispersion, cell distribution in the microchannel was analyzed (Fig. S5)† and is provided in a graph as shown in Fig. 6. To confirm the improvement of cell dispersion and performance of the device, it was compared with a reference microfluidic device which had no micropillars but had the same cell concentration and flow rate. The microscopic images and secondary processes were carried out for the analysis of the cell distribution in each droplet. The optical microscopic images are taken just before entering the orifice for final generation of microdroplets using flow-focusing technique (Fig. S5†). The cell dispersion in channels is the most important factor to determine the fraction of single cells contained in a single droplet. For a further investigation of the effects, all images from reference and pillar embedded devices were carefully analyzed to count both individual and agglomerate cells by applying a small grid (i.e. red rectangles over images, Fig. S5†). We compared the reference device and pillar incorporated device for checking the effects of microstructure. Several yellow circles (i.e. cell clusters) can be observed in the reference device while no circles presented in micropillar incorporated devices (Fig. S5†). In the reference device, less than 35% of grids contained single cells in the analyzed microchannels. This low single cell distribution became an obstacle to form uniformly sized microparticles, as well as single-cell encapsulation. On the other hand, our proposed device with pillar arrays had no clusters and 57% of grids held single cells due to continuous cell break up through cluster to pillar collision, leading to a dilution of cells over microchannels. The number of cells in each grid was
Fig. 6 The histograms are plotted by the fraction of cell containing grids.
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carefully counted and the average clearly demonstrates a significant improvement of cell dispersion as shown in Fig. 6. The comparison of generated droplet size distribution between the reference and proposed device is shown in Fig. S6.† Consequently, these data clearly support the improvement of cellular distribution in the microchannel for the single-cell encapsulation process and the uniformity of hydrogel particles. As demonstrated above, the micropillar-embedded device successfully produced a good dispersion of single cell level of E. coli in the microchannels. To confirm single-cell encapsulation, we analyzed each generated droplet right after they passed through the orifice of flow-focusing geometry. For a more accurate investigation, a high speed camera was employed and took about 10 000 frames per second for counting the total number of encapsulated cells in individual microdroplets. (Fig. 7 and S7†). The event of cell capturing happened when the dispersed cell mixture entered the flow-focusing geometry. High elongational strain separated the cells and led to the capture of single cells in each microdroplet. According to the obtained data, cell-encapsulated droplets are expressed as a graph in Fig. 7. In practical applications, the device should have reproducibility in a broad range of cell concentration. To demonstrate the feasibility of the device to work under a broad range of cell concentrations, we prepared the E. coli cells with different optical densities (OD), at 600 nm absorbance, of 0.316, 0.623, 0.641, 0.752, 0.758, 0.847, and 1.707. The overall cell concentrations are more briefly presented in Table S2† and these are much higher concentrations compared to the previous reports (i.e. 105–107 cells mL−1).28,29 After injecting the
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cells, the results for the number of cell encapsulation in each droplet were analyzed as shown in Fig. 7. The proposed device can provide a unique environment to encapsulate up to 70% single cells in the droplets, among the cell containing droplets. When using the proposed device, the fraction of droplets with less than 3 cells reached about 90% on average and a maximum of 96% can be obtained at 0.316 OD. At a relatively low cell concentration (OD ~0.8) in this study, most of the droplets contained less than 3 cells (average ~90%). On the other hand, single-cell encapsulation efficiency was dramatically decreased when the cell concentration was above OD ~0.8. Based on the obtained experimental data, it can be concluded that cell concentrations in the range of 0.3 to 0.75 OD provide a better performance to encapsulate single cells in each microdroplet. In addition, the lower encapsulation efficiency above such ranges are accounted for by a serious level of cell aggregation and optimization process issues in the flow rate of both the dispersive and continuous phases, and pillar design and dimensions. Moreover, the micropillar incorporated device can dramatically improve cell distribution in the channel under a broad range of cell concentrations as well as producing monodisperse cell encapsulated hydrogels.
Conclusions In this work, we introduced micropillars incorporated into a microfluidic device for the enhancement of aggregated cell dispersion and single-cell encapsulation using hydrogels. The effects of polymers on microbial cells were carefully investigated and the possible mechanism behind polymer-induced clusterization was proposed. In consideration of this problem, the advanced solution for splitting the clusters into not only single cells, but also to be encapsulated in hydrogel was proposed and developed using densely packed micropillar arrays. Those arrays and hydrodynamic-induced forces allowed for the continuous collision of clusters. Each dimension of the pillar arrays carefully corresponded to the cluster size in the media. This proposed approach has successfully demonstrated the possibility of this device for covering a wide range of cell concentrations. Furthermore, single E. coli cells were successfully encapsulated in individual PEG-based hydrogel microparticles and this result proved the performance of micropillars in the microchannel. Thus, the pillar arrays incorporation strategy and their unique characteristics demonstrated here can be seen to be a key component for improving cell dispersion and encapsulation efficiency for the development of cellular analysis and valuable biomaterial discovery platforms. Moreover, the combination of our results and cell-based high-throughput screening will lead to the discovery of valuable biopharmaceuticals and bio-entities from heterogeneous microorganisms.
Acknowledgements Fig. 7 (a) Fraction of number of cells among the cell containing droplets under different cell concentrations. Microscopic images of different number of cell encapsulation in each droplet corresponding to (b) one, (c) two, (d) three, and (e) more than five of cells in each droplet.
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This work was supported by BioNano Health-Guard Research Center funded by the Ministry of Science, ICT & Future Planning (MSIP) of Korea as Global Frontier Project (grant
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number H-GUARD_2013M3A6B2078945), by the Public Welfare & Safety Research Program through the National Research Foundation of Korea (NRF) funded by the MSIP of Korea (NRF-2013M3A2A1073991) and by the IT R&D program of MOTIE/KEIT [10041870] Development of Growth Management Technology for Integrating Complex Raising Building Forms of Fish and Shellfish and was also partially funded by a grant from the National Fisheries Research & Development Institute (NFRDI; contribution number RP-2013-BT-XXX).
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K. Tanizawa, A. Kondo, I. Fujii and S. Kuroda, Sci. Rep., 2013, 3, 1191–1199. K. G. Lee, T. J. Park, S. Y. Soo, K. W. Wang, B. I. Kim, J. H. Park, C. S. Lee, D. H. Kim and S. J. Lee, Biotechnol. Bioeng., 2010, 107, 747–751. A. S. Hoffman, Adv. Drug Delivery Rev., 2012, 64, 18–23. S. S. Chen, W. Fitzgerald, J. Zimmerberg, H. K. Kleinman and L. Margolis, Stem Cells, 2007, 25, 553–561. J. A. Bernate, C. Liu, L. Lagae, K. Konstantopoulos and G. Drazer, Lab Chip, 2013, 13, 1086–1092. K. K. Zeming, S. Ranjan and Y. Zhang, Nat. Commun., 2013, 4, 1625–1632. H. Amini, E. Sollier, M. Masaeli, Y. Xie, B. Ganapathysubramanian, H. A. Stone and D. D. Carlo, Nat. Commun., 2013, 4, 1826–1833. A. J. Chung, D. Pulidon, J. C. Oka, H. Amini, M. Masaeli and D. D. Carlo, Lab Chip, 2013, 3, 2942–2949. M. C. Shih, S. H. Tseng, Y. S. Weng, I. M. Chu and C. H. Liu, Sens. Actuators, B, 2013, 177, 295–300. A. D. Stroock, S. K. W. Dertinger, A. Ajdari, I. Mezic, H. A. Stone and G. M. Whitesides, Science, 2002, 295, 647–651. K. C. Tang, R. M. Wibowo, D. N. Ghista and L. Yobas, Sens. Actuators, B, 2007, 128, 340–348. K. G. Lee, J. Hong, K. W. Wang, N. S. Heo, D. H. Kim, S. Y. Lee, S. J. Lee and T. J. Park, ACS Nano, 2012, 6, 6998–7008. A. Huebner, M. Srisa-Art, D. Holt, C. Abell, F. Hollfelder, A. J. deMello and J. B. Edel, Chem. Commun., 2007, 12, 1218–1220. P. R. Marcoux, M. Dupoy, R. Mathey, A. Novelli-Rousseau, V. Heran, S. Morales, F. Rivera, P. L. Joly, J.-P. Moy and F. Mallard, Colloids Surf., A, 2011, 377, 54–62. R. M. Hernández, G. Orive, A. Murua and J. L. Pedraz, Adv. Drug Delivery Rev., 2010, 62, 711–730. T. Kuhl, Y. Guo, J. L. Alderfer, A. D. Berman, D. Leckband, J. Israelachvili and S. W. Hui, Langmuir, 1996, 12, 3003–3014. D. Marenduzzo, K. Finan and P. R. Cook, J. Cell Biol., 2006, 175, 681–686. J. N. Israelachvili, in Intermolecular and Surface Forces, Academic Press, London, 2nd edn, 1991, ch. 12, pp. 246–254. P. Jenkins and M. Snowden, Adv. Colloid Interface Sci., 1996, 68, 57–96. B. Götzelmann, R. Evans and S. Dietrich, Phys. Rev. E: Stat. Phys., Plasmas, Fluids, Relat. Interdiscip. Top., 1998, 57, 6785–6800. K. Higashitani, K. Iimura and H. Sanda, Chem. Eng. Sci., 2001, 56, 2927–2938. T. Gudipaty, M. T. Stamm, L. Sl, L. Cheung, L. Jiang and Y. Zohar, Microfluid. Nanofluid., 2011, 10, 661–669.
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Micropillar arrays enabling single microbial cell encapsulation in hydrogels† Kyun Joo Park,a Kyoung G. Lee,*b Seunghwan Seok,a Bong Gill Choi,bc Moon-Keun Lee,b Tae Jung Park,bd Jung Youn Park,e Do Hyun Kim*a and Seok Jae Lee*b Single microbial cell encapsulation in hydrogels is an important task to find valuable biological resources for human welfare. The conventional microfluidic designs are mainly targeted only for highly dispersed spherical bioparticles. Advanced structures should be taken into consideration for handling such aggregated and non-spherical microorganisms. Here, to address the challenge, we propose a new type of cylindrical-shaped micropillar array in a microfluidic device for enhancing the dispersion of cell clusters and the isolation of individual cells into individual micro-hydrogels for potential practical
Received 16th January 2014, Accepted 18th February 2014 DOI: 10.1039/c4lc00070f
applications. The incorporated micropillars act as a sieve for the breaking of Escherichia coli (E. coli) clusters into single cells in a polymer mixture. Furthermore, the combination of hydrodynamic forces and a flow-focusing technique will improve the probability of encapsulation of a single cell into each hydrogel with a broad range of cell concentrations. This proposed strategy and device would be a useful
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platform for genetically modified microorganisms for practical applications.
Introduction Cell-based technology has been employed to find valuable bio-entities from a variety of cellular species, such as animal and human cells, bacteria, yeast, insect, and plant cells, for research and industry.1–3 For practical purposes, it is essential to obtain the target cells from their heterogeneous cell population.4 As a result, cell isolation and encapsulation of single cells in 3D hydrogels have become a significant task to utilize their capability to produce biopharmaceuticals, including enzymes, antibodies, proteins, peptides, DNA, and metabolites for human welfare.5,6 So far, a great improvement in the a
Department of Chemical & Biomolecular Engineering, KAIST, 291 Daehak-ro, Yuseong-gu, Daejeon 305-701, Republic of Korea. E-mail: [email protected] b Center for Nanobio Integration & Convergence Engineering (NICE), National Nanofab Center, 291 Daehak-ro, Yuseong-gu, Daejeon 305-806, Republic of Korea. E-mail: [email protected], [email protected] c Department of Chemical Engineering, Kangwon National University, Samcheok 245-711, Republic of Korea d Department of Chemistry, Chung-Ang University, Seoul 156-756, Republic of Korea e National Fisheries Research and Development Institute, 216 Haean-ro, Gijang-up, Gijang-gun, Busan 619-705, Korea † Electronic supplementary information (ESI) available: Optical, fluorescent image of microparticles, detailed dimensions of chip designs, microscopic images of cell dispersion in LB medium, depleted polymer effect on cell aggregation, images of cell distribution in device, droplet size distribution histogram, cell captured droplet generation image, break up forces value with respect to cell aggregation size, details on the force calculation, and cell concentrations are presented. See DOI: 10.1039/c4lc00070f
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handling and isolation of single cells has been made using a combination of microfluidic systems, hydrogels, and fluorescence-activated cell sorting systems (FACSs).7,8 Those methods opened a new door to overcome traditional technical limitations, such as the requirement for external filters, and other bench-top equipment in complicated processes for high-throughput single-cell analysis and sorting.9 In microfluidic systems, techniques including T-junction droplet generation,9,10 droplet trapping arrays,5,10 flow-focusing droplet generation,11–14 direct cell printing,15 and droplet break up using microgroove structures9 have been developed. These techniques have also demonstrated their capability to isolate and encapsulate the cells for subsequent applications. Although the majority of invaluable biomaterials and biopharmaceuticals are discovered from non-spherical microbial cells, these approaches are mostly optimized for spherical cells and low cell concentration. Moreover, heterogeneous gene expression and rapid cellular growth rate lead to inevitable high cell concentrations and these factors combine to hamper the currently available high-throughput sorting systems and the utilization of their potential contribution in the discovery of useful bio-entities.16 In the material aspect, polymer-based hydrogels are the most popular materials to mimic an in vivo environment for cell culturing and provide hydrophilic, and non-toxic characteristics.12,17 It also possesses numerous pores for the easy access of media to maintain cell viability and further cultivation.18 However, polymers openly disturb the surface
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charges of cells and lead to serious cellular aggregation and sedimentation issues.19,20 Moreover, this inevitable phenomenon decreases the single level cell-encapsulation efficiency as well as producing non-uniform sizes of hydrogel particles. In consequence, a new breakthrough is required to adopt the benefits from both microfluidic systems and hydrogels for the handling of non-spherical biological entities, such as rod-shaped bacterial cells and concave disc-shaped red blood cells.21 Recently, it was reported that geometrical microstructures (i.e. pillar structures) in microfluidic devices can induce significant flow deformation and migration of microparticles or cells.22,23 Based on the addressed challenges and adopting structural benefits from previous reports, the incorporation of microstructures into a microfluidic device could be an alternative approach to separate,24,25 isolate,21 and trap26 the agglomerated microbials. Additionally, the currently developed droplet-based microfluidic system provides an ideal platform for isolation of cells in hydrogels. Herein, we develop an advanced concept of encapsulating non-spherical microbial single cells into hydrogel particles using the combination of micropillar arrays and a flow-focusing technique, by the continuous breaking up of microbial cell-clusters. For further demonstration, E. coli is used as a model of a non-spherical cell due to the ease with which its genes can be modified, a fast growth rate, and the availability of a complete library of network interactions between protein complexes.27 The different dimensions of the pillar arrays correspond to the different sizes of cell clusters, which result from the presence of polymers. The pillar arrays allow us to isolate cells from the clusters and increase the single-cell encapsulation efficiency. Of particular interest, we propose that the external force will enable the continuous break up of cell clusters, resulting in the production of more than 70% of single cells into individual hydrogel particles. By using the as-prepared microfluidic platform, we were able to demonstrate the applicability of the device for a wide range of cell concentrations for the isolation of cells and producing single-cell encapsulated hydrogels for potential practical applications.
Materials and methods Materials The SU-8 photoresist and developer solution were obtained from Microchem. The poly(ethylene glycol) diacrylate (PEGDA), potassium persulfate (KPS), D-sorbitol, N,N,N′,N′tetramethylethylenediamine (TEMED), and ethanol were purchased from Sigma-Aldrich. The polydimethylsiloxane (PDMS) (Dow Corning), Abil EM 90 (Degussa), and Grapeseed oil (G-oil, Beksul) were used as received. Microfluidic device fabrication Microfluidic devices in this research were fabricated using conventional soft lithography and a PDMS molding technique. Once the photoresist structure was successfully prepared by a typical photolithography process, it was used as the master mold for PDMS casting. The PDMS mixture was
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poured into the master mold and cured at 70 °C for 2 h. The PDMS layer was peeled off from the mold and the inlet and outlet were punched out for further connection. After the curing process, the PDMS-based microfluidic device was carefully cleaned using ethanol and deionized (DI) water to remove remaining fragments or dust from the surface of the device. The structured side of PDMS was bonded with a flat PDMS layer using a corona-discharge treatment and placed in the oven at 70 °C for 1 h. Cell preparation E. coli BL21(DE3) [F− ompT hsdSB (rB− mB−) gal dcm (DE3)] was used as a host strain, expressing an enhanced green fluorescent protein (EGFP). Recombinant E. coli cells harboring pET–6His–EGFP from a previous report17 were cultivated in a 250 mL flask containing Luria-Bertani (LB) medium supplemented with ampicillin of 50 μg mL−1 in a shaking incubator (200 rpm) at 37 °C. At an OD600 of 0.4, isopropyl-D-thiogalactopyranoside (IPTG) was added to a final concentration of 1 mM to induce the gene expression. Then, cells were further cultivated for 6 h and harvested by centrifugation. Cell growth was monitored by measuring the absorbance at 600 nm with a UV/Vis spectrophotometer (Optizen POP Nano Bio). Microdroplet generation A uniform size and shape of microdroplet was achieved using a flow-focusing method.17 The microfluidic device and other components including a tygon tube (0.2 in I.D., 0.6 in O.D.), syringe pumps, and a syringe were used in the experiment. The dispersed phase was a composition of different concentrations of E. coli in LB medium (0.5 mL), PEGDA (0.3 g), KPS (12.5 mg), D-sorbitol (4 mg), and DI water (0.5 mL). The PEGDA concentration for cell encapsulation, 23 wt%, adopted was the same as previous E. coli encapsulated hydrogel research.17 Additionally, the continuous phase was a mixture of G-oil and 2 wt% of Abil EM 90. Both the dispersed phase and the continuous phase were injected into inlets of the microfluidic device. In this study, Abil EM 90 was used as a surfactant to stabilize the microdroplets and prevent the fabricated droplet merging. The microdroplets were generated in the dripping regime and had a monodisperse size. The diameter size of the microdroplets was around 45 μm, under 3 μL min−1 of dispersed phase flow rate and 4 μL min−1 of continuous phase flow rate conditions. The generation of microdroplets was observed by a charge-coupled device camera (Eclipse Ti-S, Nikon, Japan) and carefully investigated using a high speed camera (Phantom V7.3). Polymerization of microdroplets The polymerization slowly started as soon as monomers were mixed with KPS, which is the initiator for chemical crosslinking. The generated droplets were collected in the mixture of G-oil and TEMED. In this work, TEMED chemically accelerated the free-radical crosslinking of the PEGDA. Once the polymerization was completed, the synthesized particles were centrifuged at 3 000 rpm for 20 s, followed by washing with isopropyl
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alcohol (IPA) and DI water to remove the excess amounts of Goil and other residues. The isolated particles were suspended in LB medium for further investigation (Fig. S1†).
the inlet. The final obtained microdroplets were collected in TEMED which accelerated the polymerization of PEGDA for further conversion into a hydrogel.
Results and discussion
Micropillar arrays in the channel
Design of overall experimental system
The microfluidic device is composed of three distinctive cylindrical-shaped micropillar zones and a flow-focusing region for monodisperse droplet generation. It has been reported that the cylindrical-shaped pillars disturb the flow pattern around pillars.22,23 Moreover, each geometrical dimension of the pillar arrays was selected based on the different sizes of the E. coil aggregations in the device (Table S1†). The detailed design and dimensions of the device and their corresponding scanning electron microscopy (SEM) images are illustrated in Fig. 2 and S2.† The first micropillar zone had pillar diameters of 100 μm, with a pillar gap distance of 50 μm. Compared to the first zone, the diameter of pillars in the second and third micropillar zones were 50 μm, but more densely packed, with narrow gaps of 25 and 10 μm, respectively. After passing through the entire micropillar zone which had a 600 μm width, the channel width is dramatically decreased down to 180 μm. The monodisperse microdroplets were simply generated by passing through the 50 μm orifice in the device. For an easier understanding of the micropillar array structures in the device, they were investigated using SEM. The obtained images, and the shape and dimensions of the pillars, showed that the PDMS microstructures were successfully replicated from the master mold as intended and properly incorporated in the device.
The different features of the cylindrical-shaped micropillarembedded microfluidic device were adapted to enhance the dispersion for non-spherical cells. The device and microstructures were fabricated using soft-lithography and PDMS. The device (Fig. 1a) is composed of two distinct functional regions for cell dispersion and encapsulation as follows: (1) micropillar zones for the breaking up of cell clusters (Fig. 1b) and (2) a flow-focusing droplet manipulator for encapsulating the single cells (Fig. 1c). By combining the two regions in a microfluidic device, it could consecutively break up cell-clusters into single cells and each cell could be encapsulated in microdroplets. For the hydrogel fabrication, PEGDA, as monomer, with a high concentration of the E. coli mixture and oil with surfactant were separately injected into the microfluidic device, as shown in Fig. 1. The flow rates of the PEGDA with cell mixture and oil were separately controlled by syringe pumps. The microdroplets were generated at the flow-focusing region after the PEGDA and cell mixture solution passed through three different stages of micropillar zones which were closely installed to
Polymer induced cell aggregation In the previous studies, cell aggregation may not have been critically discussed or observed because of no polymers being present in the medium and low inter-cellular interaction
Fig. 1 Schematic diagrams of the experimental system and hypothesis. (a) The reagents are injected into the microfluidic device by syringe pumps. The hypothesis schematics show the effect of (b) the micropillar arrays to split cell-clusters and (c) the encapsulation of single cells into the microdroplets using the flow-focusing technique.
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Fig. 2 A schematic illustration of the micropillar arrays in the microfluidic device and their corresponding SEM images. All scale bars are 100 μm.
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forces due to low cell concentration (105–107 cells mL−1).28,29 Recently, the combination of cell and polymer induced cell agglomeration has become critical and has needed to be solved for practical applications.30 To investigate the effect of polymers (i.e. PEGDA) on E. coli cells, the optical microscopy images of cells were taken under different polymeric concentrations and analyzed as shown in Fig. 3a–f. The small black dots in optical microscopic images are indicated as E. coli in LB media. Without the presence of polymers, most of the E. coli was properly dispersed, and no aggregation was observed, as shown in Fig. 3a and S3a.† By slowly increasing the amount of PEGDA up to 13 wt%, bacteria cells started to flocculate and form relatively small cell clusters in the media (Fig. 3b and S3b†). Furthermore, larger cell cluster formation was more clearly observed in the media under 23 wt% of PEGDA (Fig. 3c and S3c†). To understand and observe the polymer induced aggregation effects more clearly, the selected area from each image (red square boxes) was carefully processed and analyzed using image processing software, Image J. Each color and brightness difference in the 3D roughness from Fig. 3d–f implies the influence of polymers on the degree of cellular agglomeration. The bright blue color and smooth surface from the image represent highly dispersed cells in the media as shown in Fig. 3d. After introducing polymers, the surface roughness of the color profile images gradually increased and changed into dark blue first (Fig. 3e) then to purple (Fig. 3f). These transitions are mainly accounted for by the high level of cellular agglomeration in the media. The heterogeneous size of the cell clusters, in the range of 2 and 153 μm (Table S1†), disrupted the generation of monodisperse
Fig. 3 Microscopic images of cells in LB media (a) without PEGDA, (b) with 13 wt% of PEGDA, and (c) with 23 wt% of PEGDA. (d–f) Degree of aggregation analysis was done by measuring 3D roughness, which correspond to the red square boxes in each image through image processing. The moment of cell-cluster encapsulation in the microdroplets (g) before and (h) after passing though the orifice using no micropillar incorporated microfluidic device. Scale bars in (a–c) are 100 μm.
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microdroplets in the flow focusing region. Therefore, the combination of microorganism and polymers would inevitably generate non-uniform sizes of microdroplets (Fig. 3g and h), as well as hampering single-cell encapsulation for additional analysis and applications. From the observation and previous reports, the severe flocculation of cells is possibly explained in two aspects as follows: (1) the partially attached polymers on the cell surface disturb the net charge balance of the cell surface,19,20 and/or (2) the PEG-induced depletion attraction.31,32 The disruption of the net charge balance occurs as the introduced polymers in the media are affected by the net balance between the van der Waals (vdW) attraction and electrostatic repulsion, which can be understood from the Derjaguin, Landau, Verwey and Overbeek (DLVO) theory.31,33 Another possibility is the PEG-based polymers are depleted and the sudden decrease of their concentration brings an osmotic force and leads to the cellular agglomeration as illustrated in Fig. S4.†34,35 Overall, the combination of net charge disruption and depletion forces induce random cells to form a cluster, to become a seed for the sequential attachment of other co-existing cells in the same environment to minimize the energy.36
Cell cluster break up Distinct microstructures were employed for the effective dispersion of the bacteria clusters in the media. It demonstrates a sequential cellular break up through the continuous collision between micropillars and E. coli as shown in Fig. 4. As soon as the clusters were injected into the first micropillar zone, they collided with pillars and split into smaller size clusters (Fig. 4b–d). In this zone, the initial cell clusters were much bigger compared to the pillar gaps (50 μm) and separated from each other by the flow stream in the channel. These continuous collisions were repeated in the second and third micropillar zones and reduced the cluster size into single cells (Fig. 4e–f). These sequential processes for enhancing the cell dispersion can be understood from the combination of cluster collision, drag force, and elongation force. The flocculated cells resulted from relatively weak cell to cell interactions.35,37 As previously discussed, the vdW and depletion forces between cells are critically dependent on the distance of each cell. In this case, the closest neighbor cells show relatively strong depletion forces while the vdW and depletion forces between inner cells and outer cells get weaker as the size of cluster increases. This phenomenon led to continuous breaking cell clusters and consequently, it is more easily separated into single cells through providing such external forces. The repeatable breaking up phenomenon can be understood by the collapse of the force balance between the adhesion force and break up force. A detailed mechanism of this phenomenon is schematically demonstrated in Fig. 5. The observed cell break up was classified as two types of mechanism. Firstly, the drag and collision forces from the disperse phase flow were mainly accounted for by the continuous impact of
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then local laminar shear stress takes over the continuous break up. Both of these mechanisms are based on the flowing of the clusters around the micropillar arrays. Details on the force calculations are discussed and given in the ESI.†
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Micropillar arrays for single-cell encapsulation
Fig. 4 (a) Picture of the microfluidic device and (b–d) optical microscopic images of the first microstructural zone under different time lines. Photographs of further cell dispersion at (e) the second and (f) third microstructural zones. All the scale bars are 100 μm.
clusters with the micropillars along the flow direction (Fig. 5a–b). Secondly, the hydrodynamic disrupting force was another major factor for further breakup (Fig. 5c–d). The hydrodynamic force led to sequential elongation repeatedly,
Fig. 5 Schematic illustration of two break up mechanisms of (a) before and (b) after collision and elongation induced (c) before and (d) after break up.
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To investigate the influence of the cylindrical pillars on the cell dispersion, cell distribution in the microchannel was analyzed (Fig. S5)† and is provided in a graph as shown in Fig. 6. To confirm the improvement of cell dispersion and performance of the device, it was compared with a reference microfluidic device which had no micropillars but had the same cell concentration and flow rate. The microscopic images and secondary processes were carried out for the analysis of the cell distribution in each droplet. The optical microscopic images are taken just before entering the orifice for final generation of microdroplets using flow-focusing technique (Fig. S5†). The cell dispersion in channels is the most important factor to determine the fraction of single cells contained in a single droplet. For a further investigation of the effects, all images from reference and pillar embedded devices were carefully analyzed to count both individual and agglomerate cells by applying a small grid (i.e. red rectangles over images, Fig. S5†). We compared the reference device and pillar incorporated device for checking the effects of microstructure. Several yellow circles (i.e. cell clusters) can be observed in the reference device while no circles presented in micropillar incorporated devices (Fig. S5†). In the reference device, less than 35% of grids contained single cells in the analyzed microchannels. This low single cell distribution became an obstacle to form uniformly sized microparticles, as well as single-cell encapsulation. On the other hand, our proposed device with pillar arrays had no clusters and 57% of grids held single cells due to continuous cell break up through cluster to pillar collision, leading to a dilution of cells over microchannels. The number of cells in each grid was
Fig. 6 The histograms are plotted by the fraction of cell containing grids.
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carefully counted and the average clearly demonstrates a significant improvement of cell dispersion as shown in Fig. 6. The comparison of generated droplet size distribution between the reference and proposed device is shown in Fig. S6.† Consequently, these data clearly support the improvement of cellular distribution in the microchannel for the single-cell encapsulation process and the uniformity of hydrogel particles. As demonstrated above, the micropillar-embedded device successfully produced a good dispersion of single cell level of E. coli in the microchannels. To confirm single-cell encapsulation, we analyzed each generated droplet right after they passed through the orifice of flow-focusing geometry. For a more accurate investigation, a high speed camera was employed and took about 10 000 frames per second for counting the total number of encapsulated cells in individual microdroplets. (Fig. 7 and S7†). The event of cell capturing happened when the dispersed cell mixture entered the flow-focusing geometry. High elongational strain separated the cells and led to the capture of single cells in each microdroplet. According to the obtained data, cell-encapsulated droplets are expressed as a graph in Fig. 7. In practical applications, the device should have reproducibility in a broad range of cell concentration. To demonstrate the feasibility of the device to work under a broad range of cell concentrations, we prepared the E. coli cells with different optical densities (OD), at 600 nm absorbance, of 0.316, 0.623, 0.641, 0.752, 0.758, 0.847, and 1.707. The overall cell concentrations are more briefly presented in Table S2† and these are much higher concentrations compared to the previous reports (i.e. 105–107 cells mL−1).28,29 After injecting the
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cells, the results for the number of cell encapsulation in each droplet were analyzed as shown in Fig. 7. The proposed device can provide a unique environment to encapsulate up to 70% single cells in the droplets, among the cell containing droplets. When using the proposed device, the fraction of droplets with less than 3 cells reached about 90% on average and a maximum of 96% can be obtained at 0.316 OD. At a relatively low cell concentration (OD ~0.8) in this study, most of the droplets contained less than 3 cells (average ~90%). On the other hand, single-cell encapsulation efficiency was dramatically decreased when the cell concentration was above OD ~0.8. Based on the obtained experimental data, it can be concluded that cell concentrations in the range of 0.3 to 0.75 OD provide a better performance to encapsulate single cells in each microdroplet. In addition, the lower encapsulation efficiency above such ranges are accounted for by a serious level of cell aggregation and optimization process issues in the flow rate of both the dispersive and continuous phases, and pillar design and dimensions. Moreover, the micropillar incorporated device can dramatically improve cell distribution in the channel under a broad range of cell concentrations as well as producing monodisperse cell encapsulated hydrogels.
Conclusions In this work, we introduced micropillars incorporated into a microfluidic device for the enhancement of aggregated cell dispersion and single-cell encapsulation using hydrogels. The effects of polymers on microbial cells were carefully investigated and the possible mechanism behind polymer-induced clusterization was proposed. In consideration of this problem, the advanced solution for splitting the clusters into not only single cells, but also to be encapsulated in hydrogel was proposed and developed using densely packed micropillar arrays. Those arrays and hydrodynamic-induced forces allowed for the continuous collision of clusters. Each dimension of the pillar arrays carefully corresponded to the cluster size in the media. This proposed approach has successfully demonstrated the possibility of this device for covering a wide range of cell concentrations. Furthermore, single E. coli cells were successfully encapsulated in individual PEG-based hydrogel microparticles and this result proved the performance of micropillars in the microchannel. Thus, the pillar arrays incorporation strategy and their unique characteristics demonstrated here can be seen to be a key component for improving cell dispersion and encapsulation efficiency for the development of cellular analysis and valuable biomaterial discovery platforms. Moreover, the combination of our results and cell-based high-throughput screening will lead to the discovery of valuable biopharmaceuticals and bio-entities from heterogeneous microorganisms.
Acknowledgements Fig. 7 (a) Fraction of number of cells among the cell containing droplets under different cell concentrations. Microscopic images of different number of cell encapsulation in each droplet corresponding to (b) one, (c) two, (d) three, and (e) more than five of cells in each droplet.
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This work was supported by BioNano Health-Guard Research Center funded by the Ministry of Science, ICT & Future Planning (MSIP) of Korea as Global Frontier Project (grant
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number H-GUARD_2013M3A6B2078945), by the Public Welfare & Safety Research Program through the National Research Foundation of Korea (NRF) funded by the MSIP of Korea (NRF-2013M3A2A1073991) and by the IT R&D program of MOTIE/KEIT [10041870] Development of Growth Management Technology for Integrating Complex Raising Building Forms of Fish and Shellfish and was also partially funded by a grant from the National Fisheries Research & Development Institute (NFRDI; contribution number RP-2013-BT-XXX).
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