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Microfluidic-based fabrication of microgels for tissue engineering Samanipour, Roya 2016

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MICROFLUIDIC-BASED FABRICATION OF MICROGELS FORTISSUE ENGINEERINGbyRoya SamanipourBSC., (Mechanical Engineering) Chamran University, Iran. 2008MSC., (Mechanical Engineering) Tarbiat Modares University, Iran. 2012A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFMaster of ScienceinTHE COLLEGE OF GRADUATE STUDIES(Interdisciplinary Studies)THE UNIVERSITY OF BRITISH COLUMBIA(Okanagan)February 2016©Roya Samanipour, 2016The College of Graduate Studies      The undersigned certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis entitled:      Microfluidic – based Fabrication of Microgels for Tissue Engineering.                                                 Roya Samanipour submitted by   in partial fulfilment of the requirements of                                      Master of Science in Interdisciplinary Studies the degree of   .         Dr. Keekyoung Kim / Mechanical Engineering  Supervisor, Professor (please print name and faculty/school above the line)       Dr. Mina Hoorfar / Mechanical Engineering  Supervisory Committee Member, Professor (please print name and faculty/school in the line above)       Dr. Frederic Menard / Chemistry   Supervisory Committee Member, Professor (please print name and faculty/school in the line above)       Dr. Liwei Wang / Mechanical Engineering  University Examiner, Professor (please print name and faculty/school in the line above)    External Examiner, Professor (please print name and university in the line above)       January 20, 2016         (Date Submitted to Grad Studies)     Additional Committee Members include:           (please print name and faculty/school in the line above)    (please print name and faculty/school in the line above)     iiAbstractThis thesis presents the experimental and computational study of hydrogel microgelsusing flow-focusing devices. The microfluidic devices were fabricated to generatemicrogels from two immiscible phases of fluids. Conventional replica molding andphotolithography methods were used to fabricate a rectangular channel microfluidicdevice. Using the flow-focusing microfluidic devices, effects of various parameters onhydrogel pre-polymer droplet generation were investigated experimentally andcomputationally. First, three-dimensional (3D) computational simulations were conductedto study the physics of hydrogel pre-polymer droplet formation mechanism in threedifferent regimes: squeezing, dripping, and jetting regime. Subsequently, effects ofviscous, inertia and surface tension force on the gelatin methacrylate (GelMA) pre-polymer droplet generation and droplet size were studied through experiments. Finally,based on computational and experimental results, the uniformly controlled size of GelMAmicrogels was created. All experimental data were summarized by a capillary number ofthe dispersed and the continuous phases to characterize the different regimes of GelMApre-polymer droplet generation and to predict the transition of dripping to a jetting regimefor GelMA pre-polymer in the flow-focusing device. Also, two types of cells, MCF-7breast cancer cells, and 3T3 fibroblasts, were mixed in a 5 wt% GelMA pre-polymersolution used as dispersed phase. The uniform cell-laden GelMA microgels werefabricated and the cell viability was over 80%. In addition, a new method to create thepolydimethylsiloxane (PDMS) circular channel was developed using a rapid and cheap3D printing process. Due to the resolution limitation of 3D printing, the channels wereelliptical, and subsequent liquid PDMS injection process was adopted to form fullycircular channels.iiiPrefaceThe research presented in this thesis is the original work of the author. The research wasconducted under the supervision of Dr. Keekyoung Kim at the Integrated Bio-Micro/Nanotechnology Laboratory in the School of Engineering at UBC Okanagan. Partsof this thesis have been published in peer reviewed scientific journals and conferenceproceedings, and my supervisor is the co-authors of all of the publications.ContributionThe results of this thesis have been partially published in peer-reviewed journals andconferences. The detail of the author contributions is as follow,Chapter 1 is based on the published journal paper. Z. Wang*, R. Samanipour*, K.Koo, and K. Kim, “Organ-on-a-chip platforms for drug delivery and cell characterization:a review,” Sensors and Materials, vol. 27, no. 6, 487-506, 2015. I wrote half of thereview paper and Dr. Kim took care of the final edition and submission.Chapter 3 is based on the paper which is under review. R. Samanipour, Z. Wang, A.Ahmadi, and K. Kim, “Computational and experimental study of the microfluidic flow-focusing generation of hydrogel droplets,” Journal of Applied Polymer Sciences, minorrevision, 2015. I developed the numerical model of droplet generation and wrote thepaper and Dr. Kim took care of the final edition and submission.Chapter 4 is based on the paper which is under review. R. Samanipour, Z. Wang, A.Ahmadi, and K. Kim, “Computational and experimental study of the microfluidic flow-focusing generation of hydrogel droplets,” Journal of Applied Polymer Sciences, minorrevision, 2015. I have done all the lab work and data analysis and Dr. Kim took care ofthe final edition and submission.ivChapter 5 is based on the published paper. B. Parker*, R. Samanipour*, A. Ahmadi,and K. Kim, “Fabrication of circular channel microfluidic chips using post-processed 3Dprinting and PDMS injection for droplet generation,” Micro and Nano Letters, publishedonline, DOI: 10.1049/mnl.2015.0329, 2015. I have done all the lab work and Dr. Kimtook care of the final edition and submission.Article published or accepted in refereed journals (* equal contribution):1. R. Samanipour, Z. Wang, A. Ahmadi, and K. Kim, “Computational and experimentalstudy of microfluidic flow-focusing generation of hydrogel droplets,” Journal ofApplied Polymer Sciences, under review, 2015.2. R. Dai, Z. Wang, R. Samanipour, and K. Kim, “Adipose derived stem cells in tissueengineering,” Stem Cells International, DOI: 10.1155/2016/6737345, 2016.3. B. Parker*, R. Samanipour*, A. Ahmadi, and K. Kim, “Fabrication of circularchannel microfluidic chips using post-processed 3D printing and PDMS injection fordroplet generation,” Micro and Nano Letters, published online, DOI:10.1049/mnl.2015.0329, 2015.4. Z. Wang*, R. Samanipour*, and K. Kim, “The cleanroom-free rapid fabrication of aliquid conductivity sensor for surface water quality monitoring,” MicrosystemTechnologies, published online, DOI: 10.1007/s00542-015-2544-1, 2015.5. Z. Wang*, R. Samanipour*, K. Koo, and K. Kim, “Organ-on-a-chip platforms fordrug delivery and cell characterization: a review,” Sensors and Materials, vol. 27, no.6, 487-506, 2015.6. Z. Wang, R. Abdulla, B. Parker, R. Samanipour, S. Ghosh, and K. Kim, “A simpleand high-resolution stereolithography-based 3D bioprinting system using visible lightcrosslinkable bioinks,” Biofabrication, published online, DOI: 10.1088/1758-5090/7/4/045009, 2015.vOther refereed contributions:7. H. Nejad, R. Samanipour, Z. Wang, K. Kim, and M. Hoofar, “Cell-Patterning andculturing on digital microfluidics,” MicroTAS 2015, Gyeongju, Korea, Oct. 2015.8. A. Gupta, B. A. Nestor, M. D. de L. Derby, A. V. den Berg, R. Samanipour, Z.Wang, H. R. Nejad, K. Kim, M. Hoorfar, “3D cell patterning using dielectrophoresison digital microfluidics,” IEEE EMBC 2015, Milano, Italy, Aug. 2015.9. Z. Wang, R. Abdulla, B. Parker, R. Samanipour, S. Ghosh, and K. Kim, “A high-resolution stereolithography system for 3D bio-printing application,” Canada-KoreaConference on Science & Technology (CKC 2015), Calgary, Alberta, Canada, Jul.2015.10. R. Samanipour, K. Kim, “Computational and Experimental study of microfluidicflow-focusing devices for synthesizing hydrogel microtissues,” ASME  InternationalMechanical Engineering Congress & Exposition, Montreal, Quebec, Canada, Nov.2014.viTable of ContentsAbstract ......................................................................................................................... iiPreface .......................................................................................................................... iiiTable of Contents ......................................................................................................... viList of Tables ................................................................................................................ ixList of Figures................................................................................................................ xAcknowledgements...................................................................................................... xvDedication................................................................................................................... xviIntroduction............................................................................................ 1Microscale Tissue Engineering....................................................................................... 1Hydrogel Biomaterials ....................................................................................................3Microfluidic Generation of Droplets............................................................................... 5T-junction Configuration............................................................................................ 7Co-flow Glass Capillary Configuration......................................................................8Flow-focusing Configuration...................................................................................... 9Objectives of Thesis......................................................................................................11Thesis Structure ............................................................................................................ 13Microfabrication of Microfluidic Flow-Focusing Devices .................. 14Overview of Microfluidic Device ................................................................................. 14Mold Fabrication........................................................................................................... 15Spin Coating ............................................................................................................. 18Photomask Alignment and UV Exposure.................................................................19PDMS Microchannel Casting and Bonding..................................................................20Summary ....................................................................................................................... 22viiTheoretical and Computational Study of Droplet Generation ........... 23Theoretical Backgrounds .............................................................................................. 23Mode 1- Squeezing...................................................................................................26Mode 2- Dripping .....................................................................................................27Mode 3-Jetting.......................................................................................................... 27Computational Simulation ............................................................................................ 303D Modeling of Device Geometry ........................................................................... 30Numerical Model of Two Phase Flow......................................................................31Fluid Properties for Simulation ................................................................................ 33Boundary Conditions and Initial Condition.............................................................. 33Computation Simulation Results and Discussion ......................................................... 36Summary ....................................................................................................................... 40Experimental Study of Hydrogel Droplet Generation and CellEncapsulation .............................................................................................................. 41Materials ....................................................................................................................... 41Experimental Setup and Procedure ............................................................................... 43Experimental Study of Hydrogel Droplet Generation .................................................. 45Experimental Validation of Computational Simulation Results............................... 45Various Parameters for Controlling Droplet Size..................................................... 49Characterization of Hydrogel Droplet Generation Regime ......................................53Cell Encapsulation Experiments ................................................................................... 55Cell Culture............................................................................................................... 55Materials and Experimental Procedure.....................................................................56Assessment of Cell Viability .................................................................................... 58Summary ....................................................................................................................... 60Fabrication of Circular Channel Microfluidic Devices ...................... 62Overview....................................................................................................................... 62viiiMethods for Circular Microchannel Fabrication........................................................... 64Mold Fabrication Using a 3D printer........................................................................ 64PDMS Casting .......................................................................................................... 65Device Bonding ........................................................................................................ 66Liquid PDMS Injection ............................................................................................ 66Result and Discussion ...................................................................................................67Summary ....................................................................................................................... 72Conclusion and Future works.............................................................. 73Summary of Remarkable Results.................................................................................. 73Future works ................................................................................................................. 75References.................................................................................................................... 78ixList of TablesTable 2.1 Acceleration, RPM, duration, and cycles to coat SU8-2025 on a substrate ...........17Table 2.2 Baking, UV light exposure, and developing time for fabricating differentthickness of SU-8 coating on a substrate ...........................................................18Table 3.1 Material properties used in computational simulation. (Values of interfacialtension adopted from [112] [113] and they measured surface tensions by thependt drop method using a Rame-Hart model number 500-F1 advancedgoniometer). .....................................................................................................34xList of FiguresFigure 1.1 Schematic of microfluidic configurations for droplet generation. (A) T-junction microfluidic configuration. (B) Flow-focusing microfluidicconfiguration. (C) Co-flow microfluidic configuration. .......................................6Figure 2.1 Microfabrication process. (A) Fabricating microchannel mold. (B) CastingPDMS on the fabricated mold. (C) Peeling off PDMS microchannel from themold. (D) Bonding PDMS microchannel to a glass slide to build a device. .......15Figure 2.2 Photolithography process. (A) Photoresist coating. (B) Mask alignment. (C)UV exposing. (D) Developing. (F) Detail steps of photolithography process. ....17Figure 2.3 Thickness of coated SU-8 2000 versus thickness. (Adopted from [72])...............19Figure 2.4 Oxygen plasma treatment and bonding mechanism. (A) Activation of thesurfaces of PDMS and glass slide with the oxygen plasma. (B) Oxygenactivated surface with hydroxyl (-OH) group. (C) Co-valent bonding ofbetween the PDMS and glass slide. (D) Permanently bonded and sealeddevice (Red color shows hydrophilic surface for requiring further bakingtime). ................................................................................................................21Figure 2.5 Fabricated device. (A) Photographic image of testing fabricated flow-focusingdevice field with red colored liquid. (B) Microscopic image of flow-focusingcross- junction channel......................................................................................22Figure 3.1 Droplet formation of water from a faucet. (A) Formation of droplets at a lowflow rate. (B) Formation of droplets slightly increasing by the flow rate. (C)Formation of a jet stream of water at a high flow rate (Adopted from [110]). ....24Figure 3.2 Droplet formation mechanisms in the flow-focusing device. (A) Mode 1-squeezing regime at very low flow rate and very low capillary numbers ofxifluids. (B) Mode 2-dripping regime at higher flow rate and moderate capillarynumbers of fluids. (C) Mode 3-jetting regime at high capillary number offluids. ...............................................................................................................28Figure 3.3 Phase diagram and formation of dripping and jetting regime in a flow-focusingdevice  (Adopted from [83])..............................................................................29Figure 3.4 Geometry and fluids domains of the flow-focusing device simulation.................31Figure 3.5 A detail schematic of computational domain used for numerical simulation andboundary condition of the flow-focusing device................................................35Figure 3.6 Illustrations of contact angle and slip length. (A) θ is the contact angle atinterface/wall contact points and (B) β  is the slip length. ..................................35Figure 3.7 A 3D model of the computational mesh grid. ......................................................36Figure 3.8 Simulation results of the change in the GelMA droplet size. (A) Diameter ofGelMA droplets (5 wt%) under the various flow rates of continuous fluid,mineral oil with 3 wt% surfactant (Span 80). (B) Diameter of GelMAmicrogels (8 wt%) under the various flow rate of continuous fluid, mineral oilwith 3 wt% surfactant (Span 80). Flow rate of disperse phase (GelMA) was2µl/min. ............................................................................................................37Figure 3.9 Computational simulation results of droplet generation during (A) thesqueezing regime and (B) dripping regime. .......................................................39Figure 4.1 Schematic of GelMA synthesis process. .............................................................42Figure 4.2 Schematic of droplet generation in a flow-focusing device .................................44Figure 4.3 Experimental Setup ............................................................................................45Figure 4.4 Comparison of experimental results with simulation results of the change inthe GelMA droplet size. (A) Diameter of GelMA droplets (5 wt%) under thevarious flow rates of continuous fluid, mineral oil with 3 wt% surfactantxii(Span 80). (B) Diameter of GelMA microgels (8 wt%) under the various flowrate of continuous fluid, mineral oil with 3 wt% surfactant (Span 80). Flowrate of the dispersed phase (GelMA) was 2 µL/min...........................................46Figure 4.5 Representative snapshots of droplet generation during (A) the squeezingregime, (B) the dripping regime, and (C) the jetting regime. Scale bar = 100µm. ...................................................................................................................48Figure 4.6 GelMA droplet sizes versus various flow rate of oil.  (A) 5 wt% GelMA wasused for the dispersed phase and Hexadecane with different concentrations ofsurfactant was used for the continuous phase. (B) 8 wt% GelMA was used forthe dispersed phase and Hexadecane with different concentrations ofsurfactant was used for the continuous phase. (C) 5 wt% GelMA was used forthe dispersed phase and mineral oil with different concentrations of surfactantwas used for the continuous phase. (D) 8 wt% GelMA was used for thedispersed phase and mineral oil with different concentrations of surfactantwas used for the continuous phase. The number of measurement for dropletsize is five and standard deviation is shown for five droplets in each cases........50Figure 4.7 Effect of surfactant for the droplet generation. 5 wt% GelMA was used for thedispersed phase with two different solution for the continuous phase: (A)mineral oil without surfactant (B) mineral oil with 3 wt% surfactant. Flowrate of GelMA and flow rate of the continuous phase was 2 µL/min and 10µL/min, respectively. Scale bar = 100μm. .........................................................51Figure 4.8 GelMA microgels generation in flow-focusing device under various flow rateof the continuous phase, (A) Qc = 5 µL/min, (B) Qc = 10 μL/min, (C) Qc =20 μL/min, and (D) Qc = 30 μL/min. 5 wt% GelMA was used for thedispersed phase and mineral oil with 3 wt% surfactant (Span 80) was used forxiiithe continuous phase. Flow rate of dispersed phase was Qd = 2 μL/min. Scalebar = 200 μm. ...................................................................................................53Figure 4.9 A phase diagram shows three different regimes resulting from the flow-focusing device in comparison with previously reported results. .......................54Figure 4.10 Cells are cultured and confluent in flasks. (A) NIH3T3 fibroblast and (B)MCF-7 breast cancer cells show different behavior. Scale bar = 100 μm. ..........56Figure 4.11 Cell laden droplet generation (100-150 μm in diameter). 6 x 106 cells/mL ofMCF-7 breast cancer cells were mixed in 5 wt% GelMA and 2 wt% VA-086photoinitiator. Scale bar = 200 μm ....................................................................58Figure 4.12 NIH 3T3 fibroblast cells encapsulated in a microgel. (A) A phase contrastimage and (B) live/dead assayed image. Both were taken from a confocalmicroscope at day 0. Scale bar = 50 µm. ...........................................................59Figure 4.13 MCF-7 breast cancer cells encapsulated in a microgel. (A) A phase contrastimage and (B) live/dead assayed image. Both were taken from a confocalmicroscope at day 0. Scale bar = 50 µm. ...........................................................60Figure 4.14 Cell viability analysis. (n=5 microgels, Error bar: Standard Deviation, *P-value < 0.05).....................................................................................................60Figure 5.1 Device fabrication using 3D printed molds. (A) SolidWorks rendering of 3Dmodeled molds for the microfluidic chip castings with features labeled. (B)Assembled microfluidic chip with light blue dye shows channels with noleakage observable. Scale bar = 5 mm...............................................................65Figure 5.2 Optical microscope images of the cross-section of channels. (A) Cross-sectionof a channel around 200 µm in diameter prior to PDMS injection. (B) PostPDMS injection and curing procedure. Scale bar = 100 µm...............................69xivFigure 5.3 Optical microscope images of the cross-section of misaligned channels. (A)Cross-section of misaligned channel prior to PDMS injection. (B) Post PDMSinjection and curing procedure. Misaligned channel resulted in correction bythe PDMS injection. Scale bar = 100 µm. .........................................................69Figure 5.4 Scanning electron micrograph of the cross section of a circular channel..............70Figure 5.5 Junction of microfluidic chip after PDMS injection. (A) Different coloredchannels show clearly that there is no blockage of the junction during PDMSinjection and circular channel formation. Scale bar = 5 mm. (B) an opticalmicroscopic image shows the detail of the junction. Scale bar = 100 µm...........71Figure 5.6 Droplet generation from the Microfluidic chip. Scale bar=200 µm......................71Figure 6.1 Schematic of filtering microfluidic device ..........................................................76xvAcknowledgementsIn the first place, I would like to thank my supervisor, Dr. Keekyoung Kim, for hisguidance and expertise throughout graduate endeavors. He has been an invaluable sourceof inspiration and encouragement.I would also like to thank my committee members and all of my other educatorsthroughout the years whom I have learned from immensely, as well as my colleagueswhom I have enjoyed working alongside in the lab.I would like to thank my family for their continuous love and support. I am forevergrateful for their generous time and compassion.Lastly, I would like to thank my partner for being there every step along the way. Thejourney would not have been the same without you. Words cannot describe myappreciation for his endless insight.xviDedicationThis thesis is dedicated to my beloved mother, Farideh Aflaki. It is impossible to thankher adequately for everything she has done for me, from raising me to loving meunconditionally. I truly want her to know that I love her very much deep in my heart.1IntroductionTissue engineering aims to create, repair, or replace biologically functional tissues using acombination of material, mechanical, chemical and biological sciences [1]. In tissueengineering, cells isolated from the biopsied tissues are encapsulated in biodegradablematerials (e.g., hydrogels), where cells can be protected from outside environments tomaintain their cellular function. Encapsulated cells can be further proliferated inbiomimetic structures (scaffolds) which resemble cellular microenvironments. Bytransplanting the resulting construct into the patients’ body, the scaffold will be degradedover time to form a new tissue [2]. During the transplantation, cells encapsulated inhydrogel scaffolds can be prevented from outside environments. The transplantation ofcell-laden hydrogel was first used in 1964 to minimize immune rejection of transplantedcells [3]. Since then, several innovative methods have been developed to advance the areaof tissue engineering. Despite the advances, there are still some limitations to controlprecisely the cellular microenvironment [4] and fabricate vascularized complex tissuestructures [5].Microscale Tissue EngineeringMicrofabrication techniques have been developed as a powerful tool to overcome anumber of tissue engineering challenges [6][7]. The ability to engineer the size and shapeof hydrogel scaffolds using microfabrication techniques has opened new opportunities toaddress the challenges such as complex tissue architecture and vascularization [8].Microfabrication techniques, such as photolithography, micromolding, and microfluidicfabrication, have been used to fabricate micrometer-sized hydrogels for encapsulatingcells in micro structures [6][9]. In photolithography technique, a thin layer ofphotocrosslinkable hydrogel pre-polymer solution is exposed to a UV light through a2photomask. The area exposed to the UV light is polymerized and becomes a gel. Thistechnique can be used to fabricate microstructures of hydrogels in a wide range of sizesfrom micrometers to millimeters. The micromolding technique is to fabricate engineeredmicrotissue structures using a microfabricated mold. The hydrogels pre-polymer can becast and crosslinked to fabricate a variety of microstructures. Another technique tofabricate hydrogel microstructures is the microfluidic technique which is a powerfulmethod to generate well-controlled hydrogel droplets in a high-throughput way. Fluidproperties within microfluidic channels can be easily modulated and generate hydrogelswith controlled environments [6].The microfluidic technique-based cell encapsulation has been widely used in recentyears due to many advantages in comparison with other microfabrication techniques [6].The microfluidic techniques offer the ability of a high-throughput generation of well-controlled monodisperse micrometer-sized spherical or rounded microgels, as well ashollow cylindrical gel fibers that cannot be created with other microfabrication techniques[8]. The size of droplets could be easily controlled by adjusting the flow rates of fluidsand fluid properties such as viscosity and interfacial tension [10]. The microfluidictechniques also offer a high degree of control over the encapsulation process to get thedesired number of cells per particles and spatial organization [11]. Controlling dropletsize and fabricating small droplets (< 150 µm) can benefit the delivery of nutrients andoxygen to encapsulated cells in microgels. The microgels created in microfluidic devicescan be used as building blocks of tissue structures [12]. For example, microgelscontaining different types of cells are assembled into a desired 3D structure to createfunctional tissues. A successful assembly process requires precisely controlling the sizeof the microgels. The microgels can be also used as a 3D cell culturing platform to studystem cell differentiation, cell-cell interaction, and cell-microenvironment interaction3[13][14]. The study of cell co-culture and cell fate regarding neighbor cells and thesurrounding environment is also benefited from controlling the size of droplets andencapsulating various cells in each microgel [15]. The study of microenvironmentalparameters (e.g., porosity and stiffness for cancer cell migration) is very crucial in tissueengineering due to their effect on cell response and fate [16]. The cell fate is influencedby microenvironmental parameters such as interaction with other cells, exposure togrowth factors, mechanical stimuli, and shear forces [17]. Therefore, the microscale tissueengineering has many advantages over the conventional macroscale tissue engineering tomodulate precisely the cell microenvironmental parameters to enhance the controllabilityof cell behavior. This study adopted microfluidic techniques to generate microgels formicroscale tissue engineering applications.Hydrogel BiomaterialsThe properties of hydrogels used for cell encapsulation resemble extracellular matrix(ECM) in native tissues [18]. Hydrogels have been extensively used for a variety of tissueengineering applications in recent years due to their unique characteristics [19][20][21].For examples, hydrogels form hydrophilic polymeric networks that can absorb the highvolume of water without dissolving. They have native tissue-like elastic properties andbiocompatibility [18]. The porous structure of hydrogels allows cells to attach to theirmicrostructure, receive oxygen and nutrients, and release metabolic products to thesurrounding environment.There are two different types of hydrogels: synthetic (synthesized in a laboratory)and natural (obtained from natural resources) hydrogels [22]. Both synthetic and naturalhydrogels are widely used in tissue engineering applications because of theirbiocompatibility and porous structures. The natural hydrogels are generally categorizedinto two main types; protein-based hydrogel (e.g., gelatin, collagen, hyaluronic acid, and4fibrin) and polysaccharides-based hydrogel (e.g., alginate, chitosan, and agarose) [22].These natural hydrogels form networks via physical or ionic interactions. Synthetichydrogels consisted of poly (ethylene glycol), poly (acrylic acid), and poly (vinyl alcohol)are synthesized by a radical chain or step-growth polymerization reaction [23]. Differenthydrogels have different characteristics based on the polymerization methods and theirstructures which affect the cellular functions such as growth, differentiation, andmigration of encapsulated cells. Each type of hydrogels has advantages and disadvantagesin tissue engineering applications [18]. Natural hydrogels are inherently biocompatibleand resemble native ECM. Therefore, they can interact with cells easily and promotevarious cellular functions. However, special processes are required to extract naturalhydrogels from biological tissue, and the extracted volume is limited. On the contrary,synthetic hydrogels have limited cell-gel interaction in comparison to natural hydrogels.However, synthetic hydrogels are highly reproducible and readily available, and theircomposition can be controlled as needed for different cell lines.In order to encapsulate cells, the pre-polymer solutions of both synthetic and naturalhydrogels are first prepared and then crosslinked to become hydrogel scaffolds with cells.Crosslinking methods vary based on types of hydrogels. Radical polymerization can bedone by heat, light or redox reactions [24]. Photocrosslinkable hydrogels such aspolyethylene glycol and gelatin methacrylate are crosslinked by exposing them to a light.Temperature changes also crosslink hydrogels such as agarose and divalent ion crosslinkshydrogels such as alginate with Ca2+ [25]. Among them, photocrosslinkable hydrogels aremost widely used in the microscale tissue engineering with the advantage of controllingcrosslinking times. During photocrosslinking process, UV light disassociatesphotoinitiator molecules mixed with hydrogel pre-polymers into free radicals to inducethe crosslinking of hydrogel networks [24]. However, the photocrosslinking process has5the risk that unreacted free radicals can also react with cellular components such as cellmembranes, ECM proteins, and DNA [24]. This may result in low cell viability or themalignant transformation of cells exposed to UV. This risk factor can be reduced bywashing the crosslinked hydrogels immediately with PBS several times to removeunreacted photoinitiators.In the ideal tissue regeneration process, hydrogels are degraded over time and cellsreplace scaffolding structures with ECM. Therefore, degradability is particularlyimportant for hydrogels used in tissue engineering applications. In the degradationprocess, the hydrophilic backbone of polymer chains is broken down due to enzymeactivities [26][27]. The multi-functional crosslinking molecules can provide a wider rangeand tighter control over the degradation rates and mechanical stiffness of hydrogels [28].Natural hydrogels, such as collagen, gelatin, fibrin, and chitosan, are biodegradable. Also,a synthetic nondegradable hydrogels, such as polyvinyl alcohol and polyethylene glycol,can be degradable by adding ester or peptide functional groups in the crosslinking agents[28].Microfluidic Generation of DropletsAqueous monodisperse droplets are generated in a variety of microfluidic deviceconfigurations such as T-junction [29], flow-focusing [30], and co-axial [31] devices(Figure 1.1).6These microfluidic methods can generate monodisperse uniform hydrogel droplets ina high throughput way. It is also possible to fabricate two or three emulsion droplets byarranging microfluidic channels in parallel [32][33]. Based on the required dropletfrequency, droplet size, and droplet uniformity, the microfluidic devices with variousconfigurations are used for generating droplets [34]. In a microfluidic device, a hydrogelpre-polymer solution mixed with cells as a dispersed phase is entered into a central inletwhile a continuous phase fluid is entered into another inlet and surrounds the hydrogelFigure 1.1 Schematic of microfluidic configurations for droplet generation. (A) T-junctionmicrofluidic configuration. (B) Flow-focusing microfluidic configuration. (C) Co-flow microfluidicconfiguration.ABCContinuous phaseContinuous phaseContinuous phaseDispersed phaseDispersed phaseDispersed phaseContinuous phaseContinuousphase7pre-polymer solution to form droplets (Figure 1.1). Nonpolar liquids, such as mineral,fluorinated, and vegetable oil, are usually used for the continuous phase [35][36][37]. Inthe flow-focusing (Figure 1.1B) and co-flow devices (Figure 1.1C), droplets break upwhen the viscous force of the continuous phase fluid overcomes the interfacial force thatkeeps droplets connected to the disperse phase fluid. To reduce surface energy, preventcoalescence, and increase monodispersity, surfactants (e.g., span 80, tween 20) are addedto the continues phase fluid [34][15][38].T-junction ConfigurationT-junction microfluidic devices are widely used for droplet generation due to their simplefabrication steps and operation process [34]. In the T-junction configuration, thedispersed phase and continuous phase enter into two channels arranged in a perpendiculardirection as shown in Figure 1.1A. The continuous phase meets and pushes the dispersedphase at the junction. The shear force and pressure gradient generated by the continuousphase fluid make a thin neck of the dispersed phase and finally creates a droplet. The sizeof droplets depends on the flow rate and viscosity of the dispersed and continuous phasefluids, as well as the geometry of microchannels. Several researchers have used T-junction microfluidic devices to encapsulate cells in microgels and studied the behaviorof encapsulated cells. Kumachev et al. studied the effect of the elasticity of cellularmicroenvironments on cell fate by fabricating microgels with different elastic properties[39]. Tan et al. generated alginate particles with narrow size distribution using a newmethod combined with the internal gelation method [37]. In this study, the calciumcarbonate nanoparticles mixed in the alginate solution induced the internal crosslinking.Droplets formed at the T-junction were broken off by a corn oil fluid. Lectin and aceticacid in the corn oil diffused into the droplets to crosslink them. Um et al. developed theT-junction microfluidic device with three different inlets for generating cell-laden8hydrogel beads [40]. In this device, pre-polymer solution entered into the first inlet andthe oil phase containing surfactant entered into the second inlet to cut off the pre-polymersolution and generate droplets. Afterward, at the downstream of the channel, anotherliquid, which carries crosslinking materials, was injected into the third inlet to mix withthe droplets. In another study of T-junction device, alginate pre-polymer and CaCl2solutions were used as dispersed phases while hexadecane was used as a continuousphase to fabricate the alginate droplets [41]. Alginate pre-polymer, CaCl2, hexadecaneentering from three different inlets were met at the junction to form a droplet andcrosslinked by chaotic mixing in a microfluidic device.Co-flow Glass Capillary ConfigurationCo-flow devices consist of two glass capillaries−an inner glass capillary and outer glasscapillary. Two fluid phases flow in the same direction in the glass capillaries. Thedispersed phase fluid flows into the inner glass capillary. The continuous phase fluidflows into the outer capillary surrounding the inner capillary in the same direction, asshown in Figure 1.1C. Hydrogel droplets are created at the tip of the inner capillary.Glass capillary devices have advantages of inherent wettability of glass surface, chemicalresistant, and true 3D circular geometry. However, the typical fabrication method of theco-flow devices is to assemble glass capillaries manually, limiting the batch fabrication ofdevices. The quality of fabricated devices varies and depends on the person. Furthermore,co-flow glass capillary devices have a limitation of controlling the size of droplets [34].Due to the difficulty of co-flow device fabrication, few studies has been carried outusing co-flow capillary devices for droplet generation in tissue engineering application[42][43]. Sugiura et al. fabricated a micro nozzle array to control the size of alginatebeads containing cells [42][43]. In [43], the device consisted of a nozzle for alginate pre-9polymer solution and air flow channel next to the nozzle. The alginate pre-polymerdroplets were cut off by air flow, fell into CaCl2 solution. Then, alginate droplets werecrosslinked and created. The results showed that the encapsulated cells in the alginatedroplets had a higher growth rate in comparison to alginate droplets generated byconventional methods because of high diffusion efficiency.Flow-focusing ConfigurationOne of the most widely used microfluidic devices for droplet generation is flow-focusingdevices. Although more technical effort is required for the fabrication of flow-focusingdevices in comparison to T-junction, the flow-focusing devices offer bettermonodispersity and higher frequency of generating droplets up to around one thousandhertz [34]. In delicate experiments, such as cell encapsulation in microgels, flow-focusingdevices are used more than other devices [34]. Also, flow-focusing devices offer a greatflexibility in generating different size of droplets by adjusting the flow rate of bothdispersed phase and continuous phase fluids [34].Anna et al. first applied the flow-focusing geometry in a microfluidic two-phase flow[30]. In the device, dispersed phase and continuous phase fluids flowed into a cross-junction channel from different inlets, which results in generating either droplets or a jetstream. Several experimental studies have investigated the effect of different parameterson droplet generation regimes in flow-focusing devices which include the effect of theviscosity of the two phases [44][10] surface tension on droplet generation [30][45], theshape of the microfluidic channel [46][47], and the different methodologies in feeding theinflow [48].In tissue engineering, flow-focusing devices are used to fabricate droplets of abiocompatible pre-polymer solution for encapsulating cells and drugs. Researchers haveemployed flow-focusing devices to generate droplets of several types of hydrogels10including magnetic hydrogel particles [49], phenyl boronic acid groups and poly(vinylalcohol) [50], polyethylene glycol [51], gelatin methacrylate (GelMA) [52], andgrapheme oxide GelMA [53]. For the tissue engineering application, the most importantparameters are the size and type of hydrogel droplets which define the cellularmicroenvironment of the encapsulated cells [50]. Dang et.al fabricated flow-focusingdevices with three different dimensions of microchannels and investigated the effect ofdevice geometry and fluid flow rate on the generation of PEG hydrogel microparticles[51].Over the last few decades, many studies have been conducted to investigate the cellencapsulation in microgels using flow-focusing microfluidic devices. A single cell wasencapsulated into monodisperse picolitre drops using a flow-focusing device [54]. Kim etal. developed a new flow-focusing device that enhanced the cell viability by rapidlyexchanging the oil phase [55]. When cell-laden alginate was cross-linked with calcifiedoleic acid, the toxic oleic acid was transformed into a harmless mineral oil and flushedout. A 3D flow-focusing device was introduced to generate the core-shell microcapsulefor the efficient formation of cell spheroid by adding hillock in the flow-focusing device[56]. Wu et al. developed a microfluidic device integrated with fluorescence-activatedsorting to increase the single cell encapsulation rate [57]. The device consisted of twomodules−a flow-focusing module for the droplet generation and a cross-shapedhydrodynamic gating module for the droplet sorting. Capretto et al. demonstrated theformation and characterization of alginate/agarose microgels for the encapsulation ofSertoli cells [58]. The high cell viability and functionality of Sertoli cells encapsulated inalginate microgels demonstrated the effectiveness of this flow-focusing device for cellencapsulation. Köster et al. demonstrated to encapsulate, incubate, and manipulateindividual cells in hydrogel droplets [59]. This study offered greater functionality of11microfluidic cell cytometers and cell sorters, allowing assay to be performed onindividual cells in their own environment prior to sorting and analyzing. Masunaga et al.used the flow-focusing microfluidic device to encapsulate different kinds of cells incollagen microgels in the size range of 50-300 µm [60]. They cultured a variety of cells,including HepG2 cells, primary neurons, primary rat hepatocytes, NIH 3T3 mousefibroblast cells, HUVECs, and MIN6 pancreatic cells in the collagen microgels. Theresults show that cells were attached to the surface of microgels in less than 2 hours. Tostudy the proliferation of cells and the ability to create a 3D tissue construct, NIH 3T3-encapsulated microgels were placed in a PDMS mold to form a 3D tissue structure after17 hours.In addition, to experimental studies, theoretical and computational studies have beenconducted to explain the dynamics of the flow-focusing process. Jensen et al. used Stokesflow theory to analyze numerically the formation of bubbles in the flow-focusing devices[61]. Lattice Boltzmann framework was also applied to understand the dynamics of flow-focusing devices [62]. In addition to physical model-based analysis, the adaptive meshingphase field model was also used to analyze the flow-focusing process [63]. Recently,researchers numerically investigated the effect of various parameters used for generatingdroplets, such as capillary number, viscosity, and geometry [64]–[66][67]. Theoreticaland computational studies of the flow-focusing device are described in detail in Chapter4. In this thesis, the flow-focusing devices were adopted to computationally andexperimentally study the mechanism of hydrogel droplet generation.Objectives of ThesisFabricating well-controlled hydrogel droplets are crucial in tissue engineering application.Microgel droplets could either use as cell culture platforms or as building blocks of tissuefabrication. Controlling the environment of encapsulated cells is important in order to12study cell-cell interaction and cell fate, as well as desired artificial tissue fabrication. Anumber of studies have been conducted to create biocompatible microgel droplets. Thesestudies mostly focused on improving the biological and mechanical properties ofhydrogel materials. However, due to the lack of systematic study of physics to generatehydrogel droplets in the flow-focusing devices, creating uniform droplets and controllingthe size of hydrogel droplets remain challenging. Therefore, there is a critical need for asystematic study of the process to generate well-defined, cell-laden hydrogel droplet withcontrollable size. Although many studies have been conducted to investigate the effect ofdifferent parameters on water-in-oil droplet generation, these studies are not useful toapply in the hydrogel droplet generation because the fluid properties of hydrogels such asviscosity, density are much different from water. Therefore, the objectives of this thesisare1. Develop microfabrication process of flow-focusing hydrogel droplet generationdevices.2. Theoretical and numerical study of droplet generation mechanism in a flow-focusing device.3. Experimental study of droplet generation and cell-laden microgels fabrication.4. Develop circular channel flow-focusing devices for the hydrogel droplet generation.To achieve the objectives of this thesis, two different types of flow-focusing microfluidicdevices were developed. Hydrogel materials with different viscosity and surface tensionwere used for continuous and dispersed phases to study the effect of different parameterson droplet generation. As an essential part of the project, a series of numericalsimulations were carried out to study systematically the physics of droplet generation inthe fabricated microfluidic devices. Finally, optimized parameters were applied togenerate controlled hydrogel droplets with cells.13Thesis StructureChapter 1 describes a thorough literature review on tissue engineering, hydrogel materialsused in tissue engineering application, and microfluidic platform used for dropletgeneration. Chapter 2 describes fabrication methods of flow-focusing microfluidicdevices used for droplet generation. Basic microfabrication equipment and detailprocedure of device fabrication are covered in this chapter. Chapter 3 describes the basicprincipal of droplet formation in two immiscible fluids. The effect of various parameterssuch as viscosity and density of the fluid, flow rates of fluids, and device geometry ofdroplet formation is presented. This chapter also includes the numerical simulationmethod to study the droplet generation mechanism of the flow-focusing device. Thesimulation results of droplet generation for the different regime are described. Chapter 4presents the fabrication of hydrogel droplets in a flow-focusing device. This chapterillustrates materials of the dispersed phase and the continuous phase, the procedure tosynthesize hydrogels, and experimental procedure to generate hydrogel microgels. Effectof fluid properties of dispersed and a continuous phase of the droplet generation is alsodiscussed. Furthermore, the cell viability results of NIH 3T3 and MCF-7 cellsencapsulated in hydrogel droplets are presented. Chapter 5 presents the introduction of acircular channel flow-focusing device and the fabrication procedure of the device. Also,the test results of the fabricated device are described in this chapter. Finally, Chapter 6describes the contributions of this research along with suggestions for future works.14Microfabrication of Microfluidic Flow-Focusing DevicesThe microfluidic flow-focusing method has been well-established to generate dropletsfrom two immiscible phases of fluids. This method has been widely used to fabricatedroplets in a wide range of applications such as foods, cosmetics, drug delivery [68], andtissue engineering [69]. In this study, a flow-focusing microfluidic device has beendeveloped to generate monodisperse hydrogel droplets. This chapter describes thedevelopment of a microfluidic flow-focusing device using the conventionalmicrofabrication and softlithography techniques.Overview of Microfluidic DeviceThe flow-focusing microfluidic channel fabricated consisted of two inlets, one junction,one chamber, and one outlet. The masks were designed using DraftSight software(Dassault Systèmes, Vélizy-Villacoublay, France) and printed on a transparent plasticfilm known as photo masks (CAD Art Services, Inc., CA, USA). After designing themicrofluidic device, the microfluidic device was fabricated through replica moldingmethod. Figure 2.1 shows the basic steps of replica molding method used in this work.Fabrication stems consist of mold fabrication, casting, replica removal, and bonding ofreplica to a substrate to provide a sealed device. The molds were fabricated viaphotolithography and mold casting was processed through the softlithography technique.To date, the most commonly used microfabrication technology in tissue engineeringplatforms is softlithography. The main advantages of softlithography include low cost,convenient fabrication steps, the easy control of deformation, structures in micro scale,and high compatibility with a broad range of biomaterials [70][71].15The most widely used biocompatible material used in softlithography ispolydimethylsiloxane (PDMS). PDMS is gas permeable in which oxygen can be perfusedinside the channel to supply enough oxygen to the cells inside the microfluidic device[72]. PDMS is transparent from 240 – 1100 nm wavelength. It means that PDMS isoptically clear from long-wave UV to infrared radiation (IR) range, which allows severaltypes of microscope systems to capture high-quality images. Also, PDMS is an electricalinsulating elastomer with 750 kPa Young’s Modulus, which makes it very suitable forgenerating the micro deformation dynamically. In addition, PDMS is non-toxic andbiocompatible. Cells can be cultured and grown in the PDMS micro channels [72].Mold FabricationMolds to form microfluidic channels were fabricated by a photolithography techniquewhich is a well-established process for patterning microchannel structures out of aADBCFigure 2.1 Microfabrication process. (A) Fabricating microchannel mold. (B) Casting PDMS on thefabricated mold. (C) Peeling off PDMS microchannel from the mold. (D) Bonding PDMSmicrochannel to a glass slide to build a device.16photoresist on a substrate. Microfabrication process has been done in a class 100 (i.e. lessthan 100 particles in diameter of 0.5 µm or smaller per cubic foot) cleanroom facility atthe school of engineering, UBCO. To fabricate a thick, thermally, and chemically stablemicrofluidic mold, the SU-8 2025 (Microchem, MA, USA) negative photoresist wasused. SU-8 2025 is a viscous photoresist used to fabricate a thick layer of desiredstructure patterns. 20-80 µm thick photoresist coating can be achieved in a single spincoating of SU8-2025. Therefore, the photoresist was spin-coated twice to get 80-150 µmheight microfluidic channels.Figure 2.2 shows typical fabrication steps to create microfluidic channel molds. Inthe first step, a silicon wafer was washed with acetone and isopropanol three times anddried with air. Then, 1 ml of SU-8 was dispensed on the silicon substrate and spin-coatedat 500 rpm for 10 seconds with an acceleration of 100 rpm/second, and then 1500 rpm for30 seconds with the acceleration of 300 rpm/second (Table 2.1). Afterward, thephotoresist on the wafer was baked at 65°C for 5 min and then 95°C for 10 min to removethe solvent. A photomask with desired microfluidic channel patterns was then positionedon top of the wafer. A glass fixed to a vertical shoulder was then placed on the photomaskand pressed slightly to make contact between the photomask and wafer. The sample wasexposed to a UV light at 365 nm and intensity of 11 mW/cm2 for 40 seconds to create therequired patterns on the photoresist. A post exposure bake took place directly afterexposure for 10 min at 95 °C. Finally, the substrate was immersed in a SU-8 developer toremove unexposed areas of photoresist for 7 minutes. The thickness of SU-8 mold wasmeasured using a micrometer. The baking times, exposure times, and developing timesrequired for SU-8 are shown in Table 2.2. Following two sub-chapters describe detailswith regard to the spin coating, photomask alignment, and exposure.17Figure 2.2 Photolithography process. (A) Photoresist coating. (B) Mask alignment. (C) UV exposing.(D) Developing. (F) Detail steps of photolithography process.FABCDTable 2.1 Acceleration, RPM, duration, and cycles to coat SU8-2025 on a substrate18Spin CoatingSpin coating process is widely used to coat a uniform layer of photoresist on a wafer. Theprocess is to spin a wafer and spread a liquid photoresist by the acceleration of centrifugalforce. Even though the thickness of the coating depends on the viscosity andconcentration of materials, the thickness is ultimately controlled by the spin speed. Alayer of SU-8 with a thickness of 25-80 μm could be created using a different spin coatingspeed and acceleration (Figure 2.3). Different baking processes and exposure times arecorrespondingly required to the desired thickness of the SU-8 layer (Table 2.2).According to Figure 2.3, the higher spin coating speed creates, the thinner layer ofphotoresist. The maximum thickness of SU8-2025 that could be achieved is 80 μm at1000 rpm speed. To get around 100 μm thickness of SU8-2025, the spin coating processwas applied twice. SU8-2025 was spin coated on a Si wafer using a spin coating machine(Laurell Technologies Co., North Wales, PA, USA) in AMNF (Applied Micro andNanosystems Facility) at UBCO. To spin the wafer with hi-speed RPM, the Si wafer ismounted on a chuck with suction pressure to firmly hold the wafer while spinning. TheTable 2.2 Baking, UV light exposure, and developing time for fabricating different thickness of SU-8coating on a substrate19spin coating machine is also equipped with a control panel to control various spin coatingparameters.Photomask Alignment and UV ExposureA mask alignment system (Optical Associates Inc., San Jose, CA, USA) is used totransfer microchannel patterns from a photomask to the photoresist coated on the wafer.UV light attached to the mask alignment system illuminates the wafer to expose someparts of the photoresist onto UV light. The photomask is used to block some parts andprevent exposing the light pass through to the wafer. SU-8 is a negative photoresist whichmakes the exposed area cross-linked to create patterns while the unexposed area iswashed away through a developing process.The alignment system consists of a stage to hold the wafer, a stage to clasp thephotomask, and a UV illumination system. The illumination system uses established ahigh-pressure mercury plasma arc discharge lamp. This lamp emits the ultraviolet (UV)light with the wavelength of 365 nm on to the photomask with uniform irradiance [73].Figure 2.3 Thickness of coated SU-8 2000 versus thickness. (Adopted from [72])20Better resolution can be achieved when the mask is in contact with the wafer. Theduration of exposure depends on the types of photoresist and power of the UV lamp. Forthe microchannel mold fabrication in this research, the exposure time was 40 seconds.PDMS Microchannel Casting and BondingOne of softlithography techniques called a replica molding method was used to fabricatethe microchannel on a silicone elastomer which is called PDMS [74]. The microfluidicchip was fabricated by pouring the 10:1 PDMS to curing agent mixture (SYLGARD 184,Dow Corning Co., Midland, MI, USA) over the molds in a petri dish. The petri dish wasthen placed in a vacuum desiccator for approximately 90 minutes or until all the bubblesare disappeared. The mold was then placed in an oven at 70 °C for three hours to curePDMS. Finally, the PDMS with the engraved microfluidic channel was peeled off fromthe mold as shown in Figure 2.1C. Tubing holes for the inlet and outlets of the PDMSwere punched using a biopsy punch. The PDMS channel was then bonded to the glassslide. To get a perfect sealing and permanent bonding between the PDMS and a glassslide, the surfaces both PDMS and the glass slide were cleaned and dried and then ahandheld corona discharge plasma treatment machine (Electro-Technic Products,Chicago, IL, USA) was used to treat oxygen both surfaces for 5 minutes [75]. Since thecorona discharger activated the surfaces to create free oxygen, the PDMS and glass beingin contact together form a strong co-valent bonding as shown in Figure 2.4. However, thiscauses the bonding free channel area that is turned to hydrophilic due to the plasmatreatment. Since a hydrophobic channel is needed to create hydrogel droplets in oil [76], apost-bake process is required to change the surface property of the microchannel. To dothat, the microfluidic device was placed in an oven at 70oC for 5 hours for the hydroxyl (-OH) group on the channel to be eliminated to create a hydrophobic microfluidic channel.Figure 2.5 shows the fabricated flow-focusing device. The sealing of the device was21checked with the injection of a colored liquid as shown in Figure 2.5A. The detail view ofcross-junction of the flow-focusing device is shown in Figure 2.5B.Figure 2.4 Oxygen plasma treatment and bonding mechanism. (A) Activation of the surfaces ofPDMS and glass slide with the oxygen plasma. (B) Oxygen activated surface with hydroxyl (-OH)group. (C) Co-valent bonding of between the PDMS and glass slide. (D) Permanently bonded andsealed device (Red color shows hydrophilic surface for requiring further baking time).O H O H O H O H OH O H O H O H O HOH O H O HO H O H O HO H O HO H OHO O O O O O OOABCD22SummaryIn this chapter, the fabrication of the flow-focusing device using the replica moldingtechnique was described. The PDMS microchannel was fabricated by casting PDMS onthe microfabricated mold. The cured PDMS microchannel was peeled off from the moldand bonded to a glass slide to complete the flow-focusing device. To make a irreversiblebond between the PDMS and the glass slide, a handheld corona discharge plasmatreatment machine was used to treat oxygen plasma on both surfaces of the PDMS andglass. The leakage and blockage of the device were tested by injecting a colored liquidinto the channel, and no leakage was observed. The developed flow-focusing device wasused to fabricate hydrogel droplet generation for tissue engineering application.Figure 2.5 Fabricated device. (A) Photographic image of testing fabricated flow-focusing device fieldwith red colored liquid. (B) Microscopic image of flow-focusing cross- junction channel.A B 100 µm23Theoretical and Computational Study of DropletGenerationIn this chapter, theoretical backgrounds behind the droplet formation mechanism inmicrofluidic flow-focusing devices as well as the effect of different parameters on dropletsizes are described. To verify the theoretical mechanism of droplet generation, acomputational study using a finite element analysis (FEA) software was carried out.Computational study results were used to investigate parameters for hydrogel dropletgeneration experiments.Theoretical BackgroundsWhen a liquid is injected into another immiscible fluid in a microfluidic device, the liquidultimately breaks into droplets through either dripping or jetting mechanism. At a slowflow rate, the liquid drips at the orifice channel which is called a dripping regime,whereas, at a higher flow rate of flows, the liquid makes a thin stream that breaks intodroplets away from the orifice channel and this regime is called a jetting regime. Theformation of water droplets at a faucet is a good example to understand how theseregimes are created in the microfluidic devices. At a slow flow rate, water droplets arecreated close to the faucet (Figure 3.1A). In this case, surface tension causes water toform droplets at the tap, and the droplets are detached when the gravitational forceovercomes an interfacial tension force. At a higher flow rate (Figure 3.1C), the jettingregime occurs because the inertia force dominates the interface tension. At thedownstream, the jet stream breaks into droplets away from the tap because of Rayleigh-Plateau instability. Due to this theory, the jet stream of fluids was always unstable andeventually forms droplets to minimize the surface energy [31]. Any perturbation in the jetstream leads in a slightly thinner jetting. The Laplace pressure (the pressure difference24between the inside and the outside of a curved surface) within the thinner region of the jetstream increases because of the curvature of the interface.This higher pressure pushes the fluid within the jet stream to the sides, and makes the thinregion become thinner. Finally, the droplets break up once the gravitational forceovercomes the interfacial tension force. However, in the case of the microfluidic device,the droplet formation occurs due to the balance between the surface tension force and theviscous drag force of the outer liquid [77]. When a liquid is injected into anotherimmiscible liquid in a microfluidic device, the mechanism of droplet formation ischanged because of the presence of the surrounding viscous liquid [31].There are various microfluidic device geometries for the droplet generation. Amongthem, T-junction, flow-focusing, and co-flow geometries are most common in dropletgeneration (Figure 1.1). A flow-focusing microfluidic device has a well-establishedgeometry due to the advantage of fabricating high throughput monodisperse droplets. Asshown in Figure 1.1 B, in the flow-focusing device, there is an intersection of twoFigure 3.1 Droplet formation of water from a faucet. (A) Formation of droplets at a low flow rate. (B)Formation of droplets slightly increasing by the flow rate. (C) Formation of a jet stream of water at ahigh flow rate (Adopted from [117]).A B C25channels to form a cross junction. A dispersed phase fluid flows into a central channeland continuous phase fluid flows through side channels to surround the dispersed phasefluid at the junction. Then, either a jet stream or droplets of the dispersed phase fluid arecreated. The formation of either droplets or jet stream depends on different parameters,such as the geometry of devices [78][79], fluid flow rates, viscosity, density, andinterfacial energy between two fluids [80][81][10]. These parameters can be summarizedinto two non-dimensional numbers, which are called “capillary number” and “Webbernumber”. These two numbers govern the formation of droplets or jet stream in flow-focusing devices. The capillary number describes the relationship between the viscousforce and interfacial force [82] as follows,= (2.1), where μ and U represent the viscosity and velocity of fluids, respectively, and σrepresents the interfacial tension between the two immiscible fluids. The webber numberdescribes the relationship between the inertia force and the interfacial force [82] asfollows,= (2.2), where ρ, U, and D represent the density, velocity of dispersed phase fluid, and thechannel hydraulic diameter, respectively, and σ represents the interfacial tension betweenthe two immiscible fluids [83]. These nondimensional numbers are important to predictthe formation of droplets and jet stream and design experimental conditions.26To better understand the droplet generation mechanism in the flow-focusing device,three regimes of droplet formation (i.e., squeezing, dripping, and jetting regime) aredescribed in this chapter. The models developed to investigate the relationship betweenfluid properties and droplet sizes are also presented. Figure 3.2 shows three dropletgeneration modes in the flow-focusing device. In two-phase microfluidic systems,dispersed and continuous phase fluids basically are injected into the device from twoseparate microchannels. Typically, fluids meet at the junction and then the continuousphase surrounds the dispersed phase and the two-fluid interface deforms. At this point, ifthe free surface instabilities between the phases are large, droplets develop and finallybreak up from the dispersed phase. In contrast, a jet stream is formed if the free surfaceinstabilities are minimized. These instabilities come from the competition betweenstabilizing and destabilizing forces at the interface between two phases. Shear force andfluid inertia are examples of forces that promote the formation of jet stream while otherforces like capillary pressure and interfacial force promote the destabilization andcreating droplets. Three regimes of droplet formation are described in detail followingsubchapters.Mode 1- SqueezingAs shown in Figure 3.2A (Mode 1), at the very low capillary numbers of continuous anddispersed phase, the dispersed phase entering into the orifice channel blocks the channel.This results in an increase of the upstream hydrostatic pressure on the continuous phasechannel. In order to sustain the constant flow rate, the syringe pump increases thepressure applied to the continuous phase fluid streams. This leads to creating a neck ofthe dispersed phase at the entrance of orifice channel. Finally, the narrowing neckbecomes unstable and breaks to form a droplet of the dispersed phase fluid [84]. Droplets27are formed by the equilibrium of the interface tension and hydrostatic pressure field.Gastecki et al. introduced a rate-of-flow-controlled breakup model in the flow-focusingdevice at a low flow rate [84][85][82]. The rate-of-flow-controlled break up modeldescribes that the size of the droplets is only related to the ratio of flow rates of thedispersed phase and continuous phase fluids in case of a low flow rate.Mode 2- DrippingFor the dripping regime, the combination of capillary instabilities and viscous dragdefines the droplet formation in a flow-focusing device. A study demonstrated that eithercapillary instability or viscous drag alone could not induce the droplet formation [82]. Asshown in Figure 3.2B (Mode 2), by slightly increasing the flow rate of fluids, thedispersed phase becomes unstable because the surface tension force seeks to minimize theinterfacial area by creating a spherical droplet, while the viscous force of the continuousphase tries to suppress the formation of the droplet. When the shear force of thecontinuous phase fluid overcomes the interfacial surface tension, that keeps dropletsattached to the aqueous neck, the droplet breaks up [83]. Dripping regime investigatedbased on shearing model. The shearing model emphasizes that the diameter of droplets isinversely related to the capillary number [10].Mode 3-JettingThe jetting regime occurs either when the Webber number (We) of the dispersed phasefluid is high or the capillary number of the continuous phase is high. On the one hand,when the Webber number of the dispersed phase is greater than 1 due to either highdispersed phase flow rate or high density, the inertia force dominates interfacial tensionforces. This moves the aqueous neck downstream to generate the long, unstable jet streamof the disperse phase fluid. On the other hand, when the capillary number of the28continuous phase fluid is high due to either large continuous phase flow rate or highviscosity, the viscous drag force of the outer continuous phase fluid pulls the dispersedphase fluid enough to overcome the interfacial tension force. This also results in a longthin stream of the dispersed phase fluid. In  other words, if the viscous force of thecontinuous phase becomes more significant in comparison with the interfacial surfaceenergy, or if the inertial force of the dispersed phase dominates the interfacial energy, theperturbation is suppressed and results in a jetting regime [82].  This jet stream is unstabledue to instabilities and eventually breaks into droplets at the downstream of the channel.Based on Rayleigh- Plateau instability, the dispersed phase fluid becomes unstable at thedownstream of the channel because the surface tension of the dispersed phase tries tominimize the interfacial area by creating a spherical droplet. Though the viscous force ofthe dispersed phase tries to suppress droplet formation, the higher viscous force of thecontinuous phase than that of the dispersed phase finally overcomes the interfacialtension to form droplets (Figure 3.2C).Figure 3.3 shows the phase diagram of the dispersed phase and continuous phase in aflow-focusing device [86]. The figure shows the capillary numbers of dispersed phaseversus continuous phase. At the low capillary numbers, droplets are easily created in theFigure 3.2 Droplet formation mechanisms in the flow-focusing device. (A) Mode 1- squeezing regimeat very low flow rate and very low capillary numbers of fluids. (B) Mode 2-dripping regime at higherflow rate and moderate capillary numbers of fluids. (C) Mode 3-jetting regime at high capillarynumber of fluids.A. Mode 1 B. Mode 2 C. Mode 329flow-focusing device. However, a jet stream is created due to the increment of capillarynumbers by increasing either flow rate or viscosity of fluids. The transition of dripping tojetting regime occurs at the capillary number around 1.The fundamental equations of mass, momentum and energy conservation are used toinvestigate the droplet formation in a microfluidic device. Since the theoretical study ofdroplet generation requires many parameters, most of droplet generation mechanisms areanalyzed using a numerical method to investigate shear forces and pressure changes in themicrofluidic devices using separated solution domains (e.g., dispersed phase andcontinuous phase). Another important factor in the numerical simulation of dropletgeneration is to resolve the phase interface since parameters, such as density, viscosityand pressure, are not continuous between two phases.Figure 3.3 Phase diagram and formation of dripping and jetting regime in a flow-focusing device(Adopted from [86]).Jetting [44]Jetting [92]Transition [44]Dripping [44]Dripping [92]Dripping [48]Dripping [118]30Moreover, most of previous studies have investigated the droplet formation andbreakup of aqueous solutions in oil. However, the fluid dynamics of the biocompatiblepre-polymer solution is different from water, which makes optimized water-oilparameters less useful in manipulating hydrogel droplet generation. Viscosity and surfacetension are among the most important parameters influencing droplet generation regime.With the emergence of new biomaterials and hydrogels, the range of viscosity and surfacetension values in microfluidic flow-focusing devices have been extended. Therefore, it isimportant to characterize the droplet formation and breakup regimes in microfluidic flowfocusing systems for a wider range of viscosity, surface tension and capillary numberswhich have not been studied for hydrogel pre-polymer solutions. Thus, this researchutilized a numerical simulation software, COMSOL Multiphysics, to optimize the flowrate, droplet sizes, and droplet generation mechanism with various viscosities, surfacetension, and capillary numbers for hydrogel pre-polymer solutions in oil using the flow-focusing device geometry.Computational SimulationComputational simulation software (COMSOL Multiphysics 4.3b, Comsol Inc.,Burlington, MA, USA) was used to simulate the hydrogel droplet formation in a flow-focusing device. The simulations were carried out in the 3D domain to study the physicsof three different regimes in two-phase flow microchannel (dripping, squeezing, andjetting) and also study in detail how to create uniform hydrogel droplets in amicrochannel flow-focusing device.3D Modeling of Device GeometryFigure 3.4 shows the dimension of the flow-focusing device with the rectangular crosssectioned microchannel. The modeling was carried out using only a rectangular cross-section because of observations of devices. The geometry of flow-focusing devices was31defined in the global definition section of COMSOL. The simulations were carried out in3D to consider the effect of all constraints realistically. To build the 3D model of theflow-focusing device, each cross-section of inlets was created in different work planesand then extruded. A square of 90 90 µm2 was created in a XY plane and then extrudedin the distance of 300 µm. Another square of 90 90 µm2 was created in a ZY plane andthen extruded in the distance of 600 µm. Figure 3.4 also shows the modeling domain.While Fluid 1 to be dispersed into hydrogel droplets enters into the central inlet, Fluid 2(continuous phased oil) enters into both sides inlets and flow through the main channel.Numerical Model of Two Phase FlowIn this study, “Laminar Two-Phase Flow, Level Set” interface in COMSOL Multiphysics3.3b (Comsol Inc., Burlington, MA, USA) is adapted to simulate the hydrogel dropletgeneration. The laminar flow was chosen because the Reynolds number (Re = ) for allthe cases were less than 4. The fluids were assumed as Newtonian and incompressiblefluids. The level set methods, a class of numerical techniques, were first introduced byFluid 1Fluid 290μm90μmFigure 3.4 Geometry and fluids domains of the flow-focusing device simulation32Osher and Sethian and deal with fluid-interface motion that is represented implicitly. Theequation of the fluid-interface motion is numerically approximated. Level set methods areparticularly useful for problems in which the topology of the evolving interface changesduring the course of events. The fluid interface between the dispersed phase andcontinuous phase is set up by a momentum transport equation, a continuity equation, anda level-set equation [87]. These equations are as follows:∂∂t 	ρ . 	 . μ	 (2.3). 0 (2.4)∅ . ∅ . ∅ 1 ∅ ∅| ∅| 	 	 ∅ (2.5), where ρ is density (kg/ m3), u velocity (m/s), t time, µ dynamic viscosity (Pa·s), ppressure (Pa), and Fst surface tension force (N/m3). ɸ is the level set function which is inthe range of 0 to 1. The value of ɸ at the interface is considered to be 0.5 and ɸ valuesless than 0.5 corresponds to Fluid 1, whereas ɸ values greater than 0.5 corresponds toFluid 2. γ and ε are the numerical stabilization parameters. ε determines the thickness ofthe interface and it should have the same order as the computational mesh size ofelements where the interface propagates. γ determines the amount of re-initialization ofthe level set function. A suitable value for γ is the maximum value for the velocity field ofu. 1 m/s was used for the re-initialization parameter (γ) which is an approximate value of33maximum speed occurring in a fluid flow. The density and viscosity are calculated asfollows,∅ (2.6)∅ (2.7), where ρ1, ρ1, µ1, and µ1 are densities and viscosities of Fluid 1 and Fluid 2.Fluid Properties for SimulationFluid 1 entering into a center inlet is specified for hydrogel pre-polymer solution. Fluid 2entering into side inlets and flows through the main channel is specified for oil with asurfactant. Properties of Fluid 1 and 2 that used in the computational simulation, such asdensities, dynamic viscosities, and surface tension, are listed in Table 3.1. These valuesobtained from the experimental measurement. The viscosity was measured using aViscometer (Cannon Instrument Company, State College, PA, USA). Various propertiesof fluids were used to study the effect of parameters (density, viscosity, surfactant, andflow rates) on hydrogel droplet generation mechanisms and droplet size.Boundary Conditions and Initial ConditionBoundary conditions of the inlet and outlet were set to describe fluid flow conditions atthe inlets and outlet. Normal inflow velocity was defined for the inlet nodes (inlet andoutlets are shown in Figure 3.5) of the simulation model. Pressure without viscous stress(p0) was chosen for the outlet boundary condition.34Figure 3.5 shows the boundary condition for the simulation. The wetted wallboundary condition applies to all solid boundaries with the contact angle of pi/4 and sliplength equal to the mesh size parameters h. In wetted wall boundary condition, the fluid-fluid interface can move along the wall, and this boundary condition is proper for walls incontact with the fluid-fluid interface. The contact angle is the angle between the fluidinterface and the solid wall at points where the fluid interface attached to the wall. Theslip length is the distance to the position outside the wall where the extrapolatedtangential velocity component is zero (Figure 3.6).Table 3.1 Material properties used in computational simulation. (Values of interfacial tensionadopted from [119] [120] and they measured surface tensions by the pendt drop method using aRame-Hart model number 500-F1 advanced goniometer).35Figure 3.7 shows the computational mesh grid used in the simulation. A numerical meshgrid was adopted after performing grid dependence studies with different grid resolutions.Figure 3.5 A detail schematic of computational domain used for numerical simulation and boundarycondition of the flow-focusing device.Figure 3.6 Illustrations of contact angle and slip length. (A) θ is the contact angle at interface/wallcontact points and (B) β is the slip length.A B36Computation Simulation Results and Discussion3D Computational simulations were conducted with the same geometry of the fabricatedflow-focusing devices and same fluid properties of the gelatin methacrylate (GelMA)hydrogel droplet generation experiments which will be discussed in Chapter 4. Figure3.8A shows the simulation results of the change in the droplet size of 5 wt% GelMAversus the various flow rates of oil with 3wt% surfactant and Figure 3.8B shows theresults of 8wt% GelMA. The droplet size was decreased by increasing the flow rate of thecontinuous phase. The droplet size of 8 wt% GelMA, which was highly viscous, was notdecreased anymore at a flow rate greater than 30 μL/min (Figure 3.8B). The minimumsize of 8 wt% GelMA could be achieved around 40 μm when the flow rate of thedispersed phase was 2 μL/min.Figure 3.7 A 3D model of the computational mesh grid.37Figure 3.9A shows the computational simulation result of the squeezing regime. Thesimulation was conducted to visualize the physics of droplet generation during squeezingregime. The flow rate of the dispersed phase (Qd) and the continuous phase (Qc) was 2μL/min and 5 μL/min, respectively. Fluid properties of 5 wt% GelMA were used for theFigure 3.8 Simulation results of the change in the GelMA droplet size. (A) Diameter of GelMAdroplets (5 wt%) under the various flow rates of continuous fluid, mineral oil with 3 wt% surfactant(Span 80). (B) Diameter of GelMA microgels (8 wt%) under the various flow rate of continuousfluid, mineral oil with 3 wt% surfactant (Span 80). Flow rate of disperse phase (GelMA) was2µl/min.AB38dispersed phase and oil with 3 wt% surfactant (Span 80) was used for the continuousphase.Two different regimes (i.e., squeezing and dripping) was numerically simulated inthe flow-focusing geometry. In the squeezing regime, droplets were generated in theorifice channel (Figure 3.9A). As discussed in Chapter 3.1.1, this regime occurred whenthe flow rate and viscosity of fluids were low. In this regime, the dispersed phase flow(GelMA) enters into the orifice and block the channel, resulting in an increase of theupstream hydrostatic pressure (pressure at point 1 in Figure 3.9A) in order to sustain theconstant flow rate. The diameter of the droplet, in this case, was greater than the diameterof the orifice channel. The pressure changes in the microfluidic channels are shown incolor. It was found that during the squeezing regime when the dispersed phase blockedthe channel, the pressure at point 1 built up to 2 kPa. When the dispersed phase blocks thechannel in the squeezing regime the difference between before (point 1in Figure 3.9A)and after (point 2 in Figure 3.9A) cross-junction pressure was around 0.8 kPa. Thus, thedroplets break up in the squeezing regime because of the pressure build-up at the cross-junction. Garstecki et al. investigated the mechanism governing the squeezing regime in aflow-focusing device when the disperse phase is a gas [84]. Gas droplets were generatedin a flow-focusing device at low flow rates, which matches well with the results ofgenerating hydrogel droplets at low flow rates. Both studies demonstrated that dropletscan be generated at low flow rates under the squeezing regime.39During the dripping regime (shown in Figure 3.9B), the flow rate or viscosity of thecontinuous phase was much greater than the dispersed phase. As discussed in Chapter3.1.2, droplets broke because of Rayleigh-Plateau instability. In fact, the dispersed phasefluids tended to minimize its surface tension energy by forming a spherical shape.Therefore, the neck was formed by the continuous phase fluid, which dragged thedispersed phase fluid to the orifice channel. When the viscosity of the continuous phasefluid overcame the interfacial force of the disperse phase, the droplets were cut off. Thedroplets were smaller than the width of the orifice channel. In the dripping regime, thepressure does not change much before or after droplet generation. The pressure differencebetween before (point 1 in Figure 3.9B) and after (point 2 in Figure 3.9B) cross-junctionis around 0.4 kPa, which is smaller than the squeezing regime (0.8 kPa). Therefore, thedroplets break up due to the Rayleigh-Plateau instability theory.Figure 3.9 Computational simulation results of droplet generation during (A) the squeezing regimeand (B) dripping regime.AB40SummaryTheoretical background of droplet generation was discussed in this chapter. Threedifferent regimes of droplet formation in a flow-focusing device: squeezing, dripping, andjetting regime was described. Either droplets formation or jet stream depends on differentparameters, such as the geometry of devices, fluid flow rates, viscosity, density, andinterfacial energy between two fluids. To study the effect of all these parameters non-dimensional capillary numbers, which are called “capillary number” and “Webbernumber” introduced in this chapter. At a high capillary number of dispersed andcontinuous phases, a stream of the jet is created in a flow-focusing device. However, atthe capillary number below 1 the droplets are created in a flow-focusing device. To studythe mechanism of droplet formation in different regimes, a computational study usingCOMSOL software was carried out. Computational simulation results were used tooptimize parameters for experiments. The geometry of simulation, boundary condition,fluid properties, mesh grids, and simulation results was described in this chapter. Thesimulation results were comparable to the previous studies and matched well with theexperimental results of this study. Simulation results show that the droplets are formed inthe squeezing regime because of the pressure gradient in the continuous phase channelswhile the formation of droplets in a jetting regime is because of the viscous force of thecontinuous phase.41Experimental Study of Hydrogel Droplet Generation andCell EncapsulationIn this chapter, the experimental study of the GelMA droplet generation in the flow-focusing device is described. To achieve a wider range of capillary numbers, this studyquantitatively investigated the GelMA droplet generation mechanism in the flow-focusingdevice from experiments. The effects of several key parameters, such as the concentrationof hydrogel, the concentration of surfactant, and the viscosity of continuous and dispersedphases were experimentally studied. All experimental data were summarized by capillarynumbers of the dispersed phase and the continuous phase to characterize the differentregimes of droplet generation and to predict the transition of GelMA drops to jet. Takentogether, by controlling those parameters, we can control the GelMA droplet size between30 µm to 200 µm and achieve uniform droplet size. Finally, the experiment of cellencapsulation in GelMA microgels and cell viability test results is depicted.MaterialsExperiments were carried out using materials with various viscosities, densities, andconcentrations of surfactants for the continuous and dispersed phase fluids. Two differentconcentrations of GelMA pre-polymer solutions for the dispersed phase were prepared bydissolving 5 wt% and 8 wt% of GelMA in Phosphate Buffered Saline (PBS). Lightmineral oil (high viscous material) and hexadecane (low viscous material, Sigma-Aldrich,St. Louis, MO, USA) with different concentrations, 0 wt%, 3 wt%, and 20 wt% ofsurfactant (Span 80, Sigma-Aldrich, St. Louis, MO, USA) were used for the continuousphase.Gelatin methacrylate pre-polymer solution was used for the dispersed phase fluid. Toprepare the GelMA pre-polymer solution, different percentages of GelMA (5 wt%, 642wt%, and 8 wt%) were dissolved in PBS. Then, different amount of photoinitiator addedto the pre-polymer GelMA solution and then final solution placed in an oven for 30minutes at 70 o C to be fully dissolved.  Two different photoinitiators were used in thisstudy, 0.2 wt% irgacure and 2 wt% VA-086. Each photoinitiators were mixed withGelMA pre-polymer in 15 mL centrifuge tubes, and tubes placed in the oven for 30 minat 70 o C to be fully dissolved.As shown in Figure. 4.1, GelMA was synthesized by the method described in [88].Briefly, 5 g of gelatin from porcine skin (Sigma-Aldrich, St. Louis, MO, USA) wasdissolved in 45 ml of dimethyl sulfoxide (Sigma-Aldrich, St. Louis, MO, USA) at 50 oC,followed by dissolving 0.5 g of 4-(dimethylamino)-pyridine (Sigma-Aldrich, St. Louis,MO, USA) in the solution. Then, 0.5 g of glycidyl methacrylate (Sigma-Aldrich, St.Louis, MO, USA) was slowly added to the solution while it was stirring at 50 °C. Thereaction was kept at 50 °C for 48 hours. After that, the solution was dialyzed againstFigure 4.1 Schematic of GelMA synthesis process.43deionized (DI) water by using a dialysis membrane tube (molecular weight cut off:12000–14000 Da; Fisher Scientific, Waltham, MA, USA) for seven days, while thedeionized water was changed twice a day. Finally, the dried GelMA was made throughthe lyophilization process. Methacrylate group is added to make gelatinphotocrosslinkable by the reaction with photoinitiators. Gelatin (denatured protein)backbone is chemically changed so as to have methacrylate groups along the backbone.The vinyl methacrylate groups allow this polymer to react either with itself or with othervinyl monomers/macromonomers to generate permanent hydrogels. This reaction can beinitiated by UV. Gelatin itself becomes gel by cooling. However, this gel is notpermanently crosslinked so that it becomes liquid by increasing the temperature. Gelatinhas a good binding site for cell growth and thus GelMA offers a useful surface for cellattachment and growth [89].Two different oil types were used for continuous phase: light mineral oil andhexadecane. To decrease the surface tension and prevent droplets from merging, asurfactant (Span 80) with different amounts, 3 wt%, and 20 wt%, were added to themineral oil and hexadecane. Surfactants are organic materials that reduce the surfacetension (interfacial tension) between two liquids or between a liquid and a solid. They areamphiphilic, meaning they have both hydrophilic head and hydrophobic tail groups.Surfactants are adsorbed at the interface of water and oil. The water-insolublehydrophobic group extends out of the bulk water phase into the oil phase, while thewater-soluble head group remains in the water phase. Therefore, the surface energy of theinterface is reduced.Experimental Setup and ProcedureAs shown in Figure. 4.2, the flow-focusing device consists of two inlets, one outlet, ajunction, an orifice channel, and an expanding chamber. The GelMA pre-polymer44solution is injected into the main inlet while oil with a surfactant is injected into the otherinlet through syringes connected by plastic tubes (0.38 mm inner and 0.79 mm outerdiameters). Oil flowing from two opposite sides of the channel breaks GelMA pre-polymer solution at the junction, which leads to generating GelMA droplets at thejunction, the orifice channel, or the expanding chamber.Figure. 4.3 shows the experimental setup built around a microscope, syringe pumps,a heater, and a temperature control chamber. The flow rates of both GelMA and oil arecontrolled by syringe pumps (KD Scientific Inc., Holliston, MA, USA). The syringepump for injecting GelMA pre-polymer solution was placed in the temperature controlchamber as GelMA tends to become a gel at room temperature. In all experiments, theflow rate of GelMA was kept constant at 2 μL/min, and the flow rate of oil was in therange of 5-40 μL/min. For the reliability of the results of each test, the device wascontinuously operated for 5 minutes at a constant flow rate, and then experimental datawere recorded. The experiments were monitored by an inverted microscope.Figure 4.2 Schematic of droplet generation in a flow-focusing device45Experimental Study of Hydrogel Droplet GenerationExperimental Validation of Computational Simulation ResultsIn order to validate the computational results with experimental results, the droplet sizefrom experimental results was compared with simulation results with the sameparameters. Figure 4.4A shows the comparison of experimental and numerical results ofthe change in the droplet size of 5 wt% GelMA versus the various flow rates of oil with 3wt% surfactant and Figure 4.4B shows the results of 8 wt% GelMA. As shown in thesefigures, the droplet size of the simulation results matches well with the droplet size of theexperimental results (within 10% confidence interval). According to the Figure 4.4, thereis an interval at 15 µL/min fo bth 5 and 8 wt% GelMA. This interval happened because atthis flow  the regime changed from the squeezing to the dripping.Figure 4.3 Experimental SetupSyringe pumpHeaterGlass syringeMicroscopeeeMicrofluidic deviceTemperaturecontrol chamber46Three different regimes (i.e., squeezing, dripping, and jetting) of droplet generationwere demonstrated experimentally in the flow-focusing devices. Figure 4.5 showssnapshots of the droplet generation from experiments under the three regimes. In thesqueezing regime experiment, 5 wt% GelMA was used for the dispersed phase and oilwith 3 wt% surfactant (span 80) was used for the continuous phase. The flow rate of thedispersed phase (Qd) and the flow rate of the continuous phase (Qc) was 2 µL/min and 5Figure 4.4 Comparison of experimental results with simulation results of the change in the GelMAdroplet size. (A) Diameter of GelMA droplets (5 wt%) under the various flow rates of continuousfluid, mineral oil with 3 wt% surfactant (Span 80). (B) Diameter of GelMA microgels (8 wt%) underthe various flow rate of continuous fluid, mineral oil with 3 wt% surfactant (Span 80). Flow rate ofthe dispersed phase (GelMA) was 2 µL/min.AB47µL/min, respectively. As shown in Figure 4.5A, droplets were generated in the orificechannel. This regime occurred when the flow rate and viscosity of fluids were relativelylow.For the dripping regime experiment, 5 wt% GelMA was used for the dispersed phaseand oil with 3 wt% surfactant (Span 80) was used for the continuous phase. The flow rateof the dispersed phase (Qd) and the flow rate of the continuous phase (Qc) was 2 µL/minand 20 µL/min, respectively. The flow rate of the continuous phase was much greaterthan the dispersed phase. As shown in Figure 4.5B, the droplets were cut off because theviscosity of the continuous phase fluid overcame the interfacial force of the dispersedphase and the size of droplets was smaller than the width of the orifice channel. In thiscase, droplets broke because of Rayleigh-Plateau instability [90]. Both squeezing regimeand dripping regime experiment results were well matched with computational simulationresults described in Chapter 3.3.In the jetting regime (shown in Figure 4.5C), the thin stream of the dispersed phasewas created, and droplets were finally cut off in the expansion chamber as a result ofRayleigh-Plateau instability. This regime occurred when the viscosity of the continuousand dispersed phases were both high. Therefore, 8 wt% GelMA was used for thedispersed phase and oil with 20 wt% surfactant (Span 80) was used for the continuousphase. The flow rate of the dispersed phase (Qd) and flow rate of the continuous phase(Qc) was 2 µL/min and 40 µL/min respectively. The viscosity of the continuous phasedragged the dispersed phase, but the formation of the droplets was suppressed because ofthe high viscosity of the dispersed phase, resulting in the jetting regime.48Figure 4.5 Representative snapshots of droplet generation during (A) the squeezing regime, (B) thedripping regime, and (C) the jetting regime. Scale bar = 100 µm.ABC49Various Parameters for Controlling Droplet SizeIn order to study the effect of surface tension on droplet generation, hexadecane andmineral oil without surfactant were compared with hexadecane and mineral oil with 3%surfactant (Span 80). By adding surfactant up to 3% to the fluid, the surface tension of thecontinuous phase liquids was dramatically decreased. However, it did not change theviscosity of the liquids.The diameters of the droplets created by mineral oil and hexadecane, with differentpercentage of surfactant, are shown in Figure 4.6. Comparing the results of hexadecanewithout surfactant, hexadecane with 3% surfactant, mineral oil without surfactant, andmineral oil with 3% surfactant shows that when surface tension between two immisciblefluid drops, the size of droplets were decreased. In addition to the effect of surfactant ondecreasing the size of droplets, uniform monodisperse droplets were dependent on theexistence of the surfactant in the continuous phase. As shown in Figure 4.7A, dropletsthat were created without the surfactant were easily merged after generation, resulting innon-uniform droplet sizes. However, uniform monodisperse droplets were created byadding surfactants (Figure 4.7B).In order to study the effect of the dispersed phase viscosity (GelMA concentration),two different solutions of GelMA 5 wt% (Figure 4.6A and C) and 8 wt% (Figure 4.6Band D) were carried out. It was found that the generated droplet diameters were verysensitive to the concentration of GelMA. Comparing the cases with the same continuousphase, but different GelMA concentrations (as their dispersed phase), it was found thatdroplet size increases by increasing the GelMA concentration. This suggests that the sizeof the droplet does not only depend on the viscosity of a continuous phase but alsodepends on GelMA concentration. In addition, the jetting regime observed, when GelMA8 wt% and mineral oil with 20 wt% surfactant (for the dispersed phase and the continuous50phase, respectively) were used, due to the high viscosity of the continuous phase fluid anddispersed phase fluid (Figure 4.6D).Figure 4.6 GelMA droplet sizes versus various flow rate of oil.  (A) 5 wt% GelMA was used for thedispersed phase and Hexadecane with different concentrations of surfactant was used for thecontinuous phase. (B) 8 wt% GelMA was used for the dispersed phase and Hexadecane with differentconcentrations of surfactant was used for the continuous phase. (C) 5 wt% GelMA was used for thedispersed phase and mineral oil with different concentrations of surfactant was used for thecontinuous phase. (D) 8 wt% GelMA was used for the dispersed phase and mineral oil with differentconcentrations of surfactant was used for the continuous phase. The number of measurement fordroplet size is five and standard deviation is shown for five droplets in each cases.51Using a higher concentration of GelMA makes the inertia force of the dispersed phase tobe considerable, which results in creating a jetting regime. In this study, the flow rate ofhydrogel pre-polymer solution kept constant at 2 μL/min in order to prevent the formationof the jetting regime. At higher flow rates of hydrogel pre-polymer solution, the inertiaforce of dispersed phase fluid (hydrogel pre-polymer solution) becomes dominant and theWebber number of disperse phase is more than (We >1), resulting in a stream of the jet.The effect of viscosity of the continuous phase was studied by comparing cases ofmineral oil and hexadecane with 3% surfactant (µ = 4.2 mPa·s) and 20% surfactant (µ =6.7 mPa·s). Adding more than 3% surfactant to the fluid will not change the interfacialtension between the dispersed phase fluid and the continuous phase fluid because of themicelle effect [91]. However, it increases the viscosity of the fluids dramatically. Figure4.6A and B showed droplet sizes when hexadecane was used as a continuous phase.Comparing hexadecane with 3% surfactant and 20% surfactant shows that the size of thedroplets was decreased by increasing the viscosity of the continuous phase (the amount ofsurfactant). Also, the same results are shown by comparing the mineral oil andFigure 4.7 Effect of surfactant for the droplet generation. 5 wt% GelMA was used for the dispersedphase with two different solution for the continuous phase: (A) mineral oil without surfactant (B)mineral oil with 3 wt% surfactant. Flow rate of GelMA and flow rate of the continuous phase was 2µL/min and 10 µL/min, respectively. Scale bar = 100μm.52hexadecane (Figure 4.6), since the viscosity of hexadecane is much less than mineral oilwhile the surface tension of mineral oil and hexadecane is almost the same. In addition,the high viscosity of the continuous phase fluid caused the dispersed phase fluid to stretchinto long thin streams. At the high viscosity of the continuous phase, it is difficult tofabricate 8 wt% GelMA droplet. However, a higher concentration of GelMA is requiredfor fabricating stiffer microgels in tissue engineering applications. In addition, UVexposure time to crosslink the hydrogel is decreased with a higher concentration ofGelMA, resulting in higher cell viability for cell-laden hydrogel application. According toFigure 4.6, small 8 wt% GelMA droplets could be created when less viscous continuousphase fluid (e.g. hexadecane) was used. When hexadecane used as a continuous phase,due to the low viscosity of the hexadecane, GelMA droplets of two differentconcentrations (5 wt% and 8 wt%) were created through the squeezing regime. Theseresults show that hexadecane could be a good option for fabricating high concentration ofhydrogel droplets in tissue engineering application. Moreover, hexadecane presents highbiocompatibility using as a continuous phase for fabricating cell-laden alginate droplets[41].Figure 4.8 shows the effect of the flow rate for the continuous phase. The sizes of thedroplets were dramatically decreased by increasing the flow rate of a continuous phase.By increasing the flow rate of the continuous phase, the viscosity of the continuous phaseincreased, thus, the high viscosity of the continuous phase suppressed the dispersed phasefluid, resulting in the small size of droplets.53Characterization of Hydrogel Droplet Generation RegimeFigure 4.9 shows a phase diagram of GelMA droplet generation observed through theexperiments. Capillary numbers of each experiment were calculated and plotted on thephase diagram to predict the formation of GelMA droplets or a jet stream in a flow-focusing device. In flow-focusing geometry, non-dimensional capillary numbers areimportant in describing the transition between the dripping and the jetting regime,because the viscosity of the flow-focusing devices. The capillary number of thecontinuous phase Cac = μν/σ and disperse phase Cad = μν/σ described in Figure 4.9 showsthe range of formation of each regime.Figure 4.8 GelMA microgels generation in flow-focusing device under various flow rate of thecontinuous phase, (A) Qc = 5 µL/min, (B) Qc = 10 μL/min, (C) Qc = 20 μL/min, and (D) Qc = 30μL/min. 5 wt% GelMA was used for the dispersed phase and mineral oil with 3 wt% surfactant(Span 80) was used for the continuous phase. Flow rate of dispersed phase was Qd = 2 μL/min. Scalebar = 200 μm.54The result shows that at very low capillary numbers of continuous (Cac<10-2) anddispersed phase fluids (Cad<10-2), the squeezing regime happens, and droplets are createdat a size bigger than the width of the channel. When 10-2<Cac<10-1 and Cad<10-2, thedripping regime happens, and the droplets are created at a size smaller than the width ofthe channel. At high capillary numbers of continuous phase fluid (10-1 <Cac< 10) andCad>10-2, the thin jet stream of the dispersed phase fluid is created. The results show thatthe transition from the dripping regime to the jetting regime happens at Cac ~ 10-1.The capillary numbers of jetting and dripping regime observed from different studies,water in oil droplets, are also shown in the phase diagram. According to the results in [44][92] [48], most of the capillary numbers of the dispersed phase in the experiments were inthe range of Cad > 10-2 for both dripping and jetting regime. However, the results of ourexperiment shows that due to the high viscosity of hydrogels (the viscosity of 5 wt%Figure 4.9 A phase diagram shows three different regimes resulting from the flow-focusing device incomparison with previously reported results.55GelMA = 2.8 mPa·s and 8 wt% GelMA = 4.9 mPa·s), the GelMA prepolymer droplets arenot created at Cad > 10-2. At Cad > 10-2 only the jetting regime of GelMA is created.Therefore, the results from the previous studies for droplet generation of water in oilis not applicable for hydrogel droplet generation for tissue engineering application. Thisphase diagram provides a comprehensive study of hydrogel droplet generation in tissueengineering application.Cell Encapsulation ExperimentsCell CultureNIH-3T3 mouse embryonic fibroblasts were cultured in Dulbecco’s modified Eagle’smedium (Invitrogen, Carlsbad, CA, USA) supplemented with 1% penicillin-streptomycin(Invitrogen, Carlsbad, CA, USA) and 10% fetal bovine serum (Invitrogen, Carlsbad, CA,USA) at 5% CO2 and 37 °C. When cell proliferation rates reach around 80-90%confluency (Figure 4.10A), cells were passaged into new flasks. Old media was aspirated,and 5 ml of PBS was used to wash out the flask twice to remove floating dead cells. 3 mLof trypsin was used to detach the cells from the flask for 3 minutes inside the incubator.The trypsin was then deactivated with 9 mL of fresh media. The mixture of cells, trypsinand media were centrifuged at 1400 rpm for 2 minutes. The liquid phase was aspirated,and a cell pellet was suspended in 3 ml of new media. Suspended cells were seeded inflasks for culturing. The same protocol was used to culture MCF-7 breast cancer cells.However, the media used for MCF-7 cells were DMEM supplemented with 1%penicillin-streptomycin, 10% fetal bovine serum (FBS), and 1% fungizone. MCF-7 breastcancer cells cultured in a flask are shown in Figure 4.10B.56Materials and Experimental ProcedureFor a cell encapsulation experiment, two different types of cells (NIH 3T3 fibroblast andMCF-7 breast cancer cells) were detached from the flask and resuspended in the GelMApre-polymer solution. 6 x 106 cells/mL of MCF-7 cells were mixed with 5 wt% GelMApre-polymer solution and 0.5 wt% 2-Hydroxy-4′-(2-hydroxyethoxy)-2-methylpropiophenone ( Sigma-Aldrich, St. Louis, MS, USA) as a photoinitiator (PI 1).Also, 6 x 106 cells/mL of 3T3 fibroblasts were mixed with a 5 wt% GelMA pre-polymersolution and 2 wt% 2,2'-Azobis[2-methyl-N-(2-hydroxyethyl)propionamide (VA-086,Wako Pure Chemical Industries, Ltd., Japan) as another photoinitiator (PI 2). GelMA pre-polymer solution mixed with cells were used as a dispersed phase and mineral oil with 20wt% surfactant was used as a continuous phase. The flow rate of the dispersed phaseapplied to GelMA pre-polymer solution with cells was in the range of 0.3-1 μL/min andthe flow rate of continuous phase was in the range of 0.8-3 μL/min. Since adding cells inthe GelMA increased the viscosity of the solution, the low flow rate of dispersed andcontinuous phase had to apply to avoid the increment of capillary numbers.Figure 4.10 Cells are cultured and confluent in flasks (A) NIH3T3 fibroblast and (B) MCF-7 breastcancer cells show different behavior. Scale bar = 100 μm.A B57Cell-laden GelMA pre-polymer droplets were created in a flow-focusing device(Figure 4.11) and were exposed to UV light at 365 nm and intensity of 4 mW/cm2 forphotocrosslinking. The photocrosslinking process allows the spatial and temporal controlof the hydrogel formation [93]. Therefore, as the concentration of photoinitiator isincreased, the crosslinking time of GelMA is decreased. However, the amount ofphotoinitiator and UV exposure time to crosslink affect cell viability. Optimizing theminimum amount of photoinitiator and shortest crosslinking time is critical for cellviability. This study found that the minimum amount of 2-Hydroxy-4′-(2-hydroxyethoxy)-2-methylpropiophenone (PI 1) to crosslink 5 wt% GelMA was 0.5 wt%with 7 minutes UV exposure time. The minimum amount of VA-086 (PI 2) to crosslink 5wt% GelMA was 2 wt% with 5 minutes UV exposure time. Although thesephotoinitiatores crosslink the GelMA prepolymer at short wave length (254 nm) in lessthan 5 min, the GelMA prepolymer was exposed to UV at long wave length (365 nm) forlonger time in order to decrease the cell damage. Because the shortwave length of UVeffects on cell viability.The GelMA droplets containing fibroblast cells generated in the flow-focusing devicewere collected in 1.5 mL centrifuge tubes and crosslinked by UV exposure for 7 minutes.Also, GelMA droplets with MCF-7 cells were crosslinked by 5 minutes UV exposure.The surfactant and oil were separated from the cell-laden GelMA microgels using acentrifuge at 10000 rpm for 5 minutes. The centrifugation process was repeated two timesto make sure all the oil and surfactant were removed from the cell-laden GelMAmicrogels.58Assessment of Cell ViabilityTo examine the cell viability, the cell-laden microgels were stained with a Live/Dead cellviability assay (Invitrogen, Carlsbad, CA, USA). The assay consists of two fluorescencedyes,  calcein acetoxymethyl (Calcein AM) and ethidium homodimer III  (EthD-III), forstaining both live and dead cells and measures two recognized parameters of cell viabilitysuch as intercellular esterase activity and plasma membrane integrity. Calcein AM, whichis a permeable and fluorescent reagent, is converted to the intensely green fluorescentcalcein by the intracellular esterase [94]. EthD-III enters into cells through the damagedmembrane, producing a bright red fluorescence signal when binding to nucleic acids.EthD- III is excluded by the intact plasma membrane of live cells.Dye concentrations are required to adjust to achieve distinct labels of live cells withCalcein-AM and of dead cells with EthD-III. The optimal concentration of dye isdifferent for different cell types. However, it is best to use the lowest dye concentrationthat gives a sufficient signal. The standard staining solution is prepared as follows;Figure 4.11 Cell laden droplet generation (100-150 μm in diameter). 6 x 106 cells/mL of MCF-7 breastcancer cells were mixed in 5 wt% GelMA and 2 wt% VA-086 photoinitiator. Scale bar = 200 μm591- Calcein-AM and EthD-III stock solutions are removed from the freezer andallowed to warm at room temperature for 30 min.2- 4 μL of  EthD-III and 2 μL of Calcein-AM are added to 2 mL PBS.3- The stock solution is vortexed to ensure complete mixing of two dyes andkeep in a refrigerator before use.200 μL of stock solution of the live/dead assay was added to a 1 mL tube with cell-ladenGelMA microgels, and then the tube was incubated at 37 oC and 5% CO2 for 30 minutes.The cell-laden microgels in the tube were washed with PBS three times to remove theexcessive assay solution. The cell-laden microgels were examined under a fluorescentmicroscope (FLUOVIEW FV1000, Olympus, Japan) to check cell viability. Figure 4.12shows fluorescently labeled NIH 3T3 cells encapsulated in a GelMA microgels with PI 1.Figure 4.13 shows fluorescently labeled MCF-7 cells encapsulated in GelMA microgelswith PI 2. As shown in Figure 4.14, the cell viability analysis of MCF-7 (~90 %) is higherthan NIH 3T3 (~80%). The possible reason is that PI 2 with shorter UV time is morebiocompatible than that of PI 1 with longer UV time. However, a further experimentalinvestigation will be required in future works.Figure 4.12 NIH 3T3 fibroblast cells encapsulated in a microgel. (A) A phase contrast image and (B)live/dead assayed image. Both were taken from a confocal microscope at day 0. Scale bar = 50 µm.BA60SummaryThe experimental study of GelMA pre-polymer droplet generation was described in thischapter. GelMA pre-polymer solution with various concentrations was used as adispersed phase and light mineral oil and hexadecane with different amounts of surfactantwere used as a continuous phase to study the effect of various parameters on dropletgeneration in a flow-focusing device. Experiments were carried out with various flowrates of the oil and hexadecane and constant flow rate of GelMA pre-polymer solution toA BFigure 4.13 MCF-7 breast cancer cells encapsulated in a microgel. (A) A phase contrast image and (B)live/dead assayed image. Both were taken from a confocal microscope at day 0. Scale bar = 50 µm.Figure 4.14 Cell viability analysis. (n=5 microgels, Error bar: Standard Deviation, *P-value < 0.05).*61characterize the effect of continuous viscous force on the droplet size. Experimentalresults showed that the flow rates of the continuous phase controlled the size of GelMApre-polymer droplets and created squeezing, dripping, and jetting regimes. All theparameters were summarized into a non-dimensional capillary number which defines theeffect of continuous viscous force and interfacial force on GelMA pre-polymer dropletgeneration. A phase diagram of capillary numbers of the dispersed phase versuscontinuous phase was plotted to place different regimes of generating GelMA pre-polymer droplets with different conditions. The results show that the transition from thedripping regime to the jetting regime happens at Cac ~ 10-1. Finally, GelMA microgelsencapsulating two types of cells were fabricated, and the viability of cells encapsulated inGelMA microgels was tested using the Live/Dead cell viability assay. Cell viability ofhigher than 80 % was achieved in this study.62Fabrication of Circular Channel Microfluidic DevicesIn chapter 2, the fabrication method of flow-focusing microfluidic devices was described.With the conventional microfabrication method, the cross-section of the microchannel islimited to a rectangular shape. Droplet generation using the rectangular channel deviceshas drawbacks that are explained in details in the introduction of this chapter. To addressthese problems, this study developed a method to fabricate a circular microchannel flow-focusing device. This circular cross-section flow-focusing device could be used fordroplet generation to generate more uniform droplets. The fabrication process as well asresults of droplet generation using this device is presented in this chapter.OverviewA photolithography microfabrication method [95] has been used to fabricate a mastermold to cast microfluidic channel devices for droplet generation. However, the cross-sectional shape of the channels fabricated by the photolithography technique is limited toa rectangular shape. The rectangular channel has several drawbacks to generate droplets[96]. In the rectangular channel, the dispersed phase fluid wets the top and bottom wallsat a low flow rate of continuous phase fluid. Wetting walls have two disadvantages: (1)cells mixed in dispersed phase tend to attach to the walls or are damaged because of shearforce at the surface [97]; and (2) capillary instability is changed and in the result dropletbreak up is not controllable [98][99]. By increasing the flow rate of a continuous phase,the wetting surface of dispersed phase might be eliminated; however, another problemoccurs, such as device leaking and cell damage because of high shear force. Anotherimportant factor in droplet generation is the uniformity of droplets. To generate uniformdroplets, a uniform velocity profile in the cross-sectional direction is required. Since thecircular microfluidic channel is truly symmetrical unlike the rectangular cross-sectional63channel, it can generate the uniform velocity profile in the cross-sectional direction [100].Circular channels that squeeze the dispersed fluid in all direction prevent dispersed fluidto wet the walls. This results in creating well controllable, monodisperse droplets. Also,rectangular channels are more likely to clog due to aggregation of synthesized droplets[101].Due to these disadvantages, alternative fabrication methods have been developed tofabricate circular channels. For example, one method involves a trench imbedded in asilicon wafer. The trench is then filled with a thick layer of doped silicon oxide, followedby heating, which closes the trench, and creates a circular channel [102]. However, thistechnique is able only to fabricate small-sized silicon channels (a few micrometers indiameter). Another method for making circular channels utilized the capillary rise ofliquid PDMS inside an open channel in a PDMS slab [103]. This method is not practicalfor fabricating circular channels smaller than 100 μm. Abdelgawad et al. introduced anew method of fabricating circular channel by injecting liquid PDMS through arectangular channel, fabricated by the microfabrication softlithography method, followedby passing air stream in a channel filled with liquid PDMS [96]. PDMS injection fills thecorner of the rectangular channel which can make a simple circular channel. In additionto these methods, standard fabrication methods such as three dimensional (3D) printing[104][105], micro-cross construction [106] and laser machining [107] have also beenused for fabricating circular channels. Also, the circular microfluidic channel was createdusing the combination of micromolding and softlitoghraphy [108] and photolithographicreflowable photoresist [109]. The mold fabrication using these methods often requiresmany processing steps. Among these methods, due to the emergence of high resolution3D printers, 3D printing provides a simple and efficient tool for creating the molds (with64micro-sized features) that can be used for fabricating microfluidic channels with circularcross section.Recently, 3D printing has been used to fabricate circular microfluidic channels forflow focusing applications [110]. However, the fabricated circular channels were notperfectly circular. Therefore, although the resolution of 3D printing has increased,creating a perfect circular microfluidic channel for a micro droplet generation remainschallenging. In this chapter, a fast, low cost technique is presented for fabricating circularchannel flow-focusing microfluidic devices using a 3D printing method. Challengesregarding the use of 3D printed molds in the peel-off process, removing cloudy surfacefinishes in the castings, getting fully circular geometries, and fixing misalignment havebeen addressed. The flow-focusing channels are used to generate hydrogel droplets ~200µm in diameter, with the diameter of the droplet being controlled by the flow rates of thefluid [111].Methods for Circular Microchannel FabricationMold Fabrication Using a 3D printerThe mold is designed using SolidWorks® (Dassault Systems, Vélizy-Villacoublay,France) software. Two identical molds are designed with similar geometry except for thealignment pins. The male and female alignment features are made so that the two halvesof the microfluidic chip would align as desired [110]. An STL format of the mold is thenprinted (see Figure 5.1A) with a poly jet 3D printer (Object500 Connex, Stratasys Ltd.,Eden Prairie, USA) using VeroWhite-FullCure®830 (Stratasys Ltd., Eden Prairie, USA)materials at 16 µm layer resolution in Z direction and 45 µm build resolution in X-Ydirection. The molds were fabricated with a glossy surface finish to allow a smoothsurface with the roughness (Rz) of 3.8 µm previously reported in [112]. After printing,65the mold is baked at 80⁰C overnight. The part is then silanized with Trichloro(1H,1H,2H,2H-perfluro-octyl) silane (Sigma-Aldrich, St. Louis, USA). Salinization isperformed by adding 50 µL of the silane solution to a petri dish and placing the petri dishand the molds into a desiccator for 30 minutes. These processes are necessary to easilypeel the cured PDMS channel from the mold made of the VeroWhite material [113].PDMS CastingThe casting process creates the two halves of the microfluidic channel device with theirinternal geometries. It is important that the reusable molds are first properly prepared bycleaning the surface with isopropanol to ensure that the surface is particle and residuefree. The two halves of the microfluidic chip are made by pouring the liquid PDMSABFemale and MaleAlignment PinsOutlet PortCross JunctionInlet PortsFigure 5.1 Device fabrication using 3D printed molds. (A) SolidWorks rendering of 3D modeledmolds for the microfluidic chip castings with features labeled. (B) Assembled microfluidic chip withlight blue dye shows channels with no leakage observable. Scale bar = 5 mm.66(SYLGARD 184, Dow Corning Co., Midland, USA) over the molds in a petri dish. ThePDMS is made of an elastomer and a curing agent mixture with a 10:1 ratio. Once theliquid mixture is poured, it is placed in a desiccator for approximately 90 minutes or untilall the bubbles have dissipated. The mold is then placed in an oven at 70 °C for threehours to solidify the PDMS. Once the PDMS has cured, it can be peeled from the petridish and mold.Device BondingThe procedure for assembling the two halves of the microfluidic parts into a chip are asfollows.  First, the holes for the inlet and outlets are cut using a punch. To get a strongbond between the two halves of the microfluidic chip, the surfaces of both chips must beclean. To clean the PDMS parts, they are placed in an ultrasonic bath with reverseosmosis water for approximately 5 minutes. When the parts are dry, they are exposed to ahandheld corona device (BD-20, Electro-Technic Products, USA) for 5 minutes (at 30seconds/cm²) which is necessary to achieve a strong bond [75]. Finally, the two halves areput together and clamped for 72 hours to allow the bond to cure. The bonding of thedevice is checked with the injection of colored liquid shown in Figure 5.1B.Liquid PDMS InjectionTo obtain circular channels, a PDMS injection coating method [96] is adopted. A 2:1liquid heptane/'PDMS mixture is injected into the microfluidic chip, and then allowed tocure while an air stream flowed through the channels. The PDMS is the standard 10:1base to curing agent ratio. After the heptane/PDMS is fully injected into the internalgeometries of the chip, the chip is placed on a hotplate for 60 seconds at 100°C. After 60seconds, air is pumped into the chip at a constant flow rate of 24 mL/min. Air is pumpedinto all inlets and allowed to escape through all outlets. It is important that the air is67forced through all the channels to prevent blockages in the device. The chip shouldremain on the hotplate, with constant air flow, for 10 minutes. This process can berepeated to obtain smaller diameter channels with slight variations to curing times, bakingtemperatures and air flow volumes. Circular channels ranging from 200 µm to 5 µm weresuccessfully fabricated by Abdelgawad et al. [96].Result and DiscussionThe fabrication process explained in the previous section was used to fabricate a circularchannel microfluidic flow-focusing droplet generator. The half circular channels werefabricated in PDMS, and bonded to each other. If the PDMS is not fully cured (where it isin contact with the mold), it could indicate that the VeroWhite material was interferingwith the curing process. As mentioned in the previous section, this problem can bemitigated by baking and silanizing the mold. Another potential problem with casting is acloudy or rough surface finish. This can be mitigated by ensuring the mold is free of anyresidue. In addition, the curing temperature during the casting process has the potential towarp to mold. Therefore, to decrease the chance of warping the mold, it is important tokeep any temperature during casting below 70°C.As shown in Figure 5.1B, the two parts were successfully attached to each other, andno leakage was observed. Acceptable bonding strength between the two halves of thismicrofluidic chip can be difficult to obtain consistently. Thus, it is important that thesurfaces between the two halves of the microfluidic chip are in full contact with oneanother. Warped molds, or casting the PDMS on uneven surfaces, can lead to problemswith achieving full surface contact. Using a clamp is a good way to ensure full contact.The bond was found to be strong enough to carry out experiments without any leakagefrom the channels shown in Figure 5.1B.68Due to the limitations of the 3D printer, the ~200 µm channels fabricated using the3D printed molds were elliptic shaped as opposed to the desired circular. Figure 5.2Ashows a cross section of the PDMS channel after the assembly. As can be seen, the widthof the individual channel was extended and the depth of the channel was too shallow. Thechannels were elliptic shaped with pointed edges where the two halves of the PDMS jointogether. This problem was caused by the limitations of printing at the micro scale, as theprinter material cannot hold the desired half circle shape and collapses into the “mound”shape. Even 3D printers with high resolution were not able to fabricate sharp edge of thehalf circular channel. In result, a perfect circular channel was not created after bondingtwo PDMS halves. Therefore, a post PDMS injection was required to cover misalignmentand create a fully circular channel. To achieve circular channels, we injected the liquidPDMS solution and cured it by pumping air flow through the channels described in themethod section. A common problem associated with this process was the air not reachingall of the outlets. Air can create a path to one outlet, and not make a path to another. Asimple fix was to plug the outlet that has the path, and to force the air to the other outlet.Once a path has been formed by the air to all outlets, it is important that air is allowed toflow freely through all of them. If air is not allowed to flow through a channel, the PDMSwill cure, and block the entire channel. Post PDMS injection coating yields results wereshown in Figure 5.2B.69Here the channels were perfectly circular as desired. The PDMS injection coatingalso allowed us to correct misaligned channels (Figure 5.3A). This misalignment can bepresent for various reasons; however, the PDMS injection coating mitigated the problemsignificantly (Figure 5.3B). Scanning electron micrograph (SEM) of the cross section of acircular channel is shown in Figure 5.4. This figure shows that a perfect circular channelwas created after PDMS injection and all misalignments were corrected.A BFigure 5.2 Optical microscope images of the cross-section of channels. (A) Cross-section of a channelaround 200 µm in diameter prior to PDMS injection. (B) Post PDMS injection and curing procedure.Scale bar = 100 µm.A BFigure 5.3 Optical microscope images of the cross-section of misaligned channels. (A) Cross-sectionof misaligned channel prior to PDMS injection. (B) Post PDMS injection and curing procedure.Misaligned channel resulted in correction by the PDMS injection. Scale bar = 100 µm.70In this microfluidic chip, two different phases of liquids are pumped into the centerchannel (water phase) and the outer channels (oil phase). The fluids meet at the cross-junction where the viscous force of the continuous oil phase flow overcomes the surfacetension of water phase flow, which causes water droplets to be created. The cross-junction is a critical portion of this flow-focusing microfluidic chip. Figure 5.5demonstrated that the cross-junction was not affected by liquid PDMS coating to formcircular channels. As shown in Figure 5.6, the droplet generation was successful, andfluid flowed smoothly with no trace of blockage or leakage. Hydrogel droplets ofdifferent size were achieved through this process.Figure 5.4 Scanning electron micrograph of the cross section of a circular channel.71The 3D printed mold is a relatively simple manufacturing process to fabricatemicrofluidic channel devices. The 3D printing technique has advantages to rapidlyfabricate complex structures with high aspect ratio comparing with the conventionalphotolithography technique which requires photomasks and a series of microfabricationprocess. Although the current resolution of 3D printers has a limitation to fabricatestructures with hundreds of micrometers, in combination with the PDMS injectiontechnique, 3D printed molds will be a promising solution to provide the flexibility toA BFigure 5.5 Junction of microfluidic chip after PDMS injection. (A) Different colored channels showclearly that there is no blockage of the junction during PDMS injection and circular channelformation. Scale bar = 5 mm. (B) an optical microscopic image shows the detail of the junction. Scalebar = 100 µmFigure 5.6 Droplet generation from the Microfluidic chip. Scale bar=200 µm72fabricate various sizes of circular channels using the replica-molding softlithographytechnique. Therefore, the developed fabrication method using 3D printed molds canovercome the size limitation of microfluidic channel and will facilitate to use the 3Dprinting technique for fabricating a variety of microfluidic channel devices. For futurework, this rapid fabricated device will be used to encapsulate cells to generate injectablemicroscale tissues and has many potential applications in biomedical engineering area.SummaryIn this chapter, a simple and low cost fabrication method is proposed for microfluidicdroplet generators with circular channels. The PDMS channels were successfully curedand peeled off the 3D printed molds, and challenges regarding the interference of the 3Dprinted material as well as the cloudy (and rough 3D printed) surface finishes wereaddressed. Moreover, to create fully circular channels, and address challenges regardingthe misalignment of the upper and lower halves, a PDMS injection coating method wasutilized and optimized. The channels were successfully fabricated, and a flow-focusingmicrofluidic device for creating microgel droplets was assembled. No leakage wasobserved, and due to the use of circular channels, the device successfully created dropletsof different size without blockage. The developed methodology provides a simple andeffective way of fabricating circular microfluidic channels for numerous biological andchemical applications.73Conclusion and Future worksSummary of Remarkable ResultsMicrofluidics flow-focusing devices are a promising platform for generating microgelsthat can be widely used in various applications such as tissue engineering, drug delivery,and cosmetic. In the tissue engineering application, cells are encapsulated in themicrogels to produce injectable microscale tissues and study the effect ofmicroenvironment on cell behaviors and cell-cell interactions. Investigating the effect ofvarious parameters on droplet generation and optimizing these parameters are importantto fabricate well-controlled cell-laden microgels.In this thesis project, microfluidic flow-focusing devices were designed andfabricated. The conventional replica molding method using a microfabricated mold wasadopted to fabricate the flow-focusing device. The physics of hydrogel pre-polymerdroplet generation mechanism was computationally studied in three different regimes(i.e., squeezing, dripping, and jetting) and compared with previous results mostlydiscussing water droplet generation. In a squeezing regime, the droplets are usuallycreated due to hydrodynamic pressure, while the dripping regime is due to the viscousdrag force of continuous phase and the jetting regime is due to Rayleigh-Plateauinstability. With the computational simulation, it was found that the concentration ofGelMA dramatically affected the viscosity of pre-polymer solution and creation ofdroplets. GelMA pre-polymer droplets created with GelMA 8 wt% are bigger thanGelMA 5 wt% at the same flow rate. The jetting regime observed due to the highviscosity and density of dispersed phase fluid, when GelMA 8 wt% was used as thedispersed phase at high flow rate of the continuous phase. It was found that increasing theflow rate ratio of the continuous phase and dispersed phase resulted in creating smallerdroplets.74Using the fabricated flow-focusing devices, this study also experimentallyinvestigated the effect of fluid properties, including viscosity, density, surface tension,and the flow rate of the dispersed and continuous phase for generating hydrogel pre-polymer droplets. GelMA pre-polymer solution with two different concentrations, 5 wt%,and 8 wt%, were used as the dispersed phase. Two different oils (i.e., hexadecane andlight mineral oil) with 0 wt%, 5 wt%, and 20 wt% surfactants were used as the continuousphase. Experiments were conducted using different flow rates of continuous phase tostudy the influence of effective forces on droplet generation and optimize the hydrogeldroplet generation. Three different regimes were investigated experimentally andcompared with computational simulation results. All the results were summarized in twonone dimensional numbers, Cac, and Cad, to determine the range of the squeezing,dripping, and jetting regimes in a flow-focusing device for hydrogel droplets and shownin the phase diagram.The presented phase diagram demonstrated that the range of capillary numberscreated different regimes for generating GelMA pre-polymer droplets. It was found that athigher capillary numbers of GelMA pre-polymer solution droplets cannot be createdbecause of the viscosity of the pre-polymer solution and resulted in the jetting regime,while water droplets could be created in the same condition based on the result ofprevious researches. Therefor, the phase diagram of water in the oil droplet generationfrom previous researchers is not helpful for hydrogel pre-polymer droplet generationbecause of the difference in viscosity and density between hydrogel and water.The optimized parameters for GelMA pre-polymer droplet generation were used toencapsulate cells in GelMA microgels. By controlling the flow rate of the two phases, thetype of oil, and the concentration of surfactant, the droplet size of the phase with differentGelMA concentrations can be manipulated between 30 µm to 250 µm, which will be very75useful to control further the number of encapsulated cells and their surroundingmicroenvironment. MCF-7 breast cancer cells and 3T3 fibroblasts were encapsulated inthe GelMA microgels and high cell viability >80%  demonstrated that the flow-focusingsystem for generating GelMA pre-polymer droplets was a promising platform to fabricatecell-laden microgels for tissue engineering applications.In addition to the microfabricated rectangular channel devices, a new method tofabricate circular channels has been introduced since the microfabrication method for themold fabrication has limitations to fabricate only rectangular shape channels. Withadvantages of generating more uniform droplets using circular channels due to theuniform velocity profile produced in the circular channel cross section, the new methodhas been developed using 3D printed molds for upper and lower halves. To create fullycircular channels, and address challenges regarding the misalignment of the upper andlower halves, a PDMS injection coating method was utilized and optimized. The channelswere successfully fabricated, and the device generated hydrogel pre-polymer droplets.The developed methodology provides a simple and effective way of fabricating circularmicrofluidic channels for numerous biological and chemical applications.Future worksThere are several future works that are suggested in this thesis to improve the flow-focusing droplet generation system. Improve the method of separating the GelMA microgels from the continuousphase fluid (oil and surfactant): This study used a centrifugation method to filteroil out from the microgels. The process required several repetitions ofcentrifugation and re-suspending the microgels in PBS solution, resulting in losingmany microgels during the process. For the future work, an addition of filteringchamber to the flow-focusing devices will help solve the problem of losing76microgels during the oil filtration process. The proposed design of filteringchamber is given in Figure 6.1. There are two inlets and two outlets. In the inlet 2,the microgel droplets with oil are injected into the filtering chamber, while PBS isinjected into the inlet 1 with a higher flow rate to wash the oil out on themicrogels. Then the microgels with PBS are coming out from the outlet 1, whilethe oil and surfactant are coming out from the outlet 2. Improve cell viability: Using the UV light with low intensity, the crosslinkingtakes longer than 5 min which may cause harmful effects on cells. Our labrecently developed an ultrafast photocrosslinking method (< 1 min) using a laserdiode. The laser-based method can be also applied to reduce the crosslinking timeand increase the cell viability further.Even distribution of cells: Evenly distributing cells in each droplet is currently difficultsince cells tend to aggregate over time. Accordingly, cell numbers in droplets created inthe flow focusing device were not controlled. For the future work, a microfluidic mixercan be added to the system to mix evenly cells with the hydrogel pre-polymer solutionduring the entire experimental process. A magnet mixer can be also used to mix the cellsInlet 1Inlet 2Outlet 1Outlet 2Figure 6.1 Schematic of filtering microfluidic device77in a syringe right before injecting into the tubing. Our lab is planning to place a smallmagnetic stirring bar inside the syringe and we will be able to mix the cells evenly bystring the magnetic bar using a magnet rotor. In addition, a dielectrophoresis method canbe also adapted to sort out droplets with and without cells [114][115][116].78References[1] R.Lanza, R. Langer, J. P. Vacanti, eds. “Principles of tissue engineering.” AcademicPress., 2000.[2] N. Pallua, Ch. Suschek, eds. " Tissue Engineering: From lab to clinic.". SpringerScience & Business Media, 2010.[3] T. M. Chang, “Semipermeable Microcapsules.,” Science, vol. 146, no. 3643, pp.524–525, 1964.[4] A. Khademhosseini, R. 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