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Hydrogel-based microfluidic system for cell culture Chen, Michael C.W. 2009

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Hydrogel-Based Microfluidic System for Cell Culture by  Michael C.W. Chen B.Sc., Electrical and Computer Engineering, University of Waterloo, Canada, 2005 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF APPLIED SCIENCE in THE FACULTY OF GRADUATE STUDIES (Biomedical Engineering)  The University of British Columbia (Vancouver) April 2009 ©Michael C.W. Chen, 2009  Abstract Traditionally, cell culture has been done in culture flasks or well plates where the volumes and length scales involved in the culture environment are many orders of magnitude larger than the size scale of individual cells. To better tailor medical care to an individual patient, it may be necessary to carry out genetic, physiological, and biochemical analyses on very small cell samples and to have an in vitro cell culture environment that more closely approximates the in vivo conditions. A microfluidic device that integrates both cell handling and long-term 3-D cell culture techniques is presented. The designed microdevice traps cells with alginate, an ionically cross-linking hydrogel, which mimics the extra cellular matrix within our body. To encapsulate the cells, a solution of calcium ions is introduced in parallel with the alginate precursor cell suspension. Alginate hydrogel forms at the interface and as the region of gel grows it traps cells inside. This is a reversible process; the gel matrix can be dissolved and the cells can be released by the addition of ethylene-diaminetetraacetic acid (EDTA), a calcium chelator. To show that the microfluidic device is reliable for long term mammalian cell culture, hepatocytes and breast tumor cells were cultured within the alginate gel layer inside the microfluidic device for more than two weeks. Hepatocytes were able to form three-dimensional aggregates within the microfluidic hydrogel environment. We further demonstrate the possibility of performing anticancer agent screening within this device. Breast tumor cells seeded in the microchannel were treated with doxorubicin, a common chemotherapy drug. Compared to controls, the doxorubicin inhibited cell proliferation. ii  In future, this system will have applications in cell-based testing and in studies involving small cell populations, such as cancer cells obtained from needle biopsies of tumors.  iii  Contents Abstract ............................................................................................................................... ii Contents ............................................................................................................................. iv List of Tables ................................................................................................................... viii List of Figures .................................................................................................................... ix Acknowledgments............................................................................................................. xii Dedication ........................................................................................................................ xiii CHAPTER 1: Introduction ................................................................................................ 1 1.1  From Semiconductors to Biomedical Microdevices........................................... 1  1.2  Micro Total-Analysis System (µ-TAS) .............................................................. 2  1.3  Cell Trapping in Microfluidic Chip .................................................................... 4  1.3.1  Chemical Trapping...................................................................................... 5  1.3.2  Hydrodynamic Trapping............................................................................. 5  1.3.3  Dielectrophoretic (DEP) Trapping.............................................................. 6  1.3.4  Optical Trapping ......................................................................................... 6  1.3.5  Acoustic Trapping....................................................................................... 7  1.3.6  Magnetic Trapping...................................................................................... 7  1.3.7  Hydrogel Trapping...................................................................................... 8  1.4  Traditional Cell Culture and 3-D Cell Culture in Microchannel ........................ 8  1.4.1  Traditional Cell Culture .............................................................................. 8  1.4.2  3-D Cell Culture in Microchannel .............................................................. 9  1.4.3  Co-culture Multiple Cell Types ................................................................ 10 iv  1.5  Motivation of this Work.................................................................................... 11  CHAPTER 2: Alginate Hydrogel Cell Encapsulation ..................................................... 14 2.1  Applications of Hydrogel.................................................................................. 14  2.2  Cell Encapsulation and Cultivation .................................................................. 15  2.2.1  Naturally Derived Hydrogel ..................................................................... 16  2.2.1.1  Collagen ................................................................................................ 16  2.2.1.2  Agarose ................................................................................................. 16  2.2.1.3  Alginate (Alginic Acid) ........................................................................ 17  2.2.2  Synthetic Hydrogel ................................................................................... 17  2.2.2.1  Poloxamer ............................................................................................. 18  2.2.2.2  N-isopropylacrylamide Polymers (NiPAAM) ...................................... 19  2.3  Chemical Structure of Alginate Hydrogel ........................................................ 19  2.4  Mechanical Properties of LF120M Sodium Alginate....................................... 22  2.5  Alginate Gel Stability ....................................................................................... 24  CHAPTER 3: Microfabrication Technology ................................................................... 26 3.1  Polydimethlsiloxane (PDMS) Microfluidic Chip ............................................. 26  3.1.1  Chemical Structure and Property of PDMS.............................................. 27  3.1.2  PDMS Curing............................................................................................ 27  3.1.3  Diffusivity of PDMS Chip ........................................................................ 29  3.1.3.1  Oxygen Supply...................................................................................... 29  3.1.3.2  Nutrient Supply..................................................................................... 30  3.2 3.2.1  PDMS Bonding................................................................................................. 31 Reversible PDMS Bonding....................................................................... 31  v  3.2.2 3.3  Irreversible PDMS Bonding ..................................................................... 31  SU-8 Photolithography ..................................................................................... 33  CHAPTER 4: Microfluidic Chip Design......................................................................... 37 4.1  Drag and Laminar Flows .................................................................................. 37  4.2  Cell Trapping .................................................................................................... 39  4.3  Diffusion and Gel Growth ................................................................................ 41  4.3.1  Gel Formation ........................................................................................... 41  4.3.2  Gel Dissolving .......................................................................................... 44  4.4  Microfluidic Chip Design Realization .............................................................. 46  4.4.1  Chip and Mask Design.............................................................................. 46  4.4.2  Micro-Fabrication Process ........................................................................ 47  4.4.2.1  SU-8 Master Mould .............................................................................. 47  4.4.2.2  PDMS Channel ..................................................................................... 48  4.4.3  Polyethyleneimine Coating (PEI) ............................................................. 51  CHAPTER 5: On-Chip Cell Culture................................................................................ 55 5.1  Biocompatibility Test........................................................................................ 55  5.1.1  Food Dye Coloring ................................................................................... 55  5.1.2  PDMS Curing............................................................................................ 57  5.2  Experimental Setup........................................................................................... 60  5.2.1  Cell Counting ............................................................................................ 61  5.2.2  Pressure-driven Fluid Flow Control System............................................. 63  5.2.3  Nutrient Circulation .................................................................................. 65  5.2.4  Live/Dead Assay....................................................................................... 69  vi  5.2.4.1  Trypan Blue .......................................................................................... 69  5.2.4.2  Live/Dead Viability/Cytotoxicity Kit ................................................... 70  5.3  Cell Viability..................................................................................................... 71  5.3.1  HFL1 Fibroblast Cells............................................................................... 71  5.3.2  Hep G2 Hypatocyte Cells ......................................................................... 73  5.3.3  LCC6\HER2 Breast Cancer Cells............................................................. 77  5.4  Co-culture ......................................................................................................... 80  5.5  Cell Releasing ................................................................................................... 82  5.6  Drug Testing ..................................................................................................... 84  5.6.1  Doxorubicin (Dox).................................................................................... 85  5.6.2  Drug Diffusion in Alginate Gel ................................................................ 86  5.6.3  Drug Diffusion through PDMS Chip........................................................ 89  5.6.4  Control Group ........................................................................................... 91  5.6.5  Experimental Result.................................................................................. 93  CHAPTER 6: Conclusion ................................................................................................ 97 6.1  Summary ........................................................................................................... 97  6.2  Future Work ...................................................................................................... 98  Bibliography ................................................................................................................... 100 Appendices...................................................................................................................... 111 A.  Status of Cell Culture Inside Microdevices ............................................................ 111  B.  Chip Fabrication Recipe ......................................................................................... 116  C.  Mammalian Cell Suspension Preparation Protocol ................................................ 117  D.  Drug Testing with LCC6/Her2 and Doxorubicin ................................................... 118  vii  1 List of Tables Table 4.1-1 Reynolds Number………………………….. …………………………....  38  Table 5.3-1 Cell proliferation after 72 hours treatment...………………………...…...  96  viii  2 List of Figures Figure 1.2.1: Micro Total-Analysis System.……….…...………………..…………...  3  Figure 1.3.1: Cell-immobilization methods……………………………………….….  4  Figure 2.2.1: Different cellular structures in 2-D and 3-D scaffolds [48]…………….  15  Figure 2.3.1: Monomers of alginic acid.....………………………………………...…  20  Figure 2.3.2: Molecular structure of alginic acid……...………………..….…………  20  Figure 2.3.3: Bonding sites of Ca2+ ion.………………………………………………  21  Figure 2.4.1: The viscosity of LF120M alginate solutions…………………………...  23  Figure 2.5.1: Stability of alginate in microchannel…….………...…………………...  25  Figure 3.1.1: The backbone structure of PDMS……………………………………...  27  Figure 3.1.2: PDMS curing………………………………………….………………..  28  Figure 3.2.1: PDMS-PDMS bonding.……… ……………………………..…………  32  Figure 3.3.1: Molecular structure of SU-8......………………………………………..  34  Figure 4.1.1: Laminar flow in the microfluidic chip.……………….……………..….  39  Figure 4.2.1: Operation principle.………………………………..…………………...  40  Figure 4.3.1: Gel formation….……………………………….…….……………..…..  42  Figure 4.3.2: Alginate gel growth rate.………………………………….……………  44  Figure 4.3.3: Alginate gel dissolving rate.....…………………………………………  45  Figure 4.4.1: Channel design……………………………………...………………….  47  Figure 4.4.2: The fabrication process of the microfluidic chip……………………….  50  Figure 4.4.3: The finished microfluidic chip…….…………………………………...  51  ix  Figure 4.4.4: Molecular structure of PEI......…………………………………………  52  Figure 4.4.5: Excess PEI solution cause alginate solution to polymerize……………  53  Figure 4.4.6: The static contact angle measurement of PEI coated PDMS surface…..  54  Figure 5.1.1: Experimental setup of food dye compatibility test ..……….…..………  56  Figure 5.1.2: Cell viability reading………………..…...…………………..…………  57  Figure 5.1.3: Experimental setup of PDMS curing compatibility test…..……..….….  59  Figure 5.1.4: Cell viability reading over four-day period……………........……..…...  60  Figure 5.2.1: Hemocytometer………………………….………….………………….  61  Figure 5.2.2: Number of trapped cells vs. initial cell suspension concentration……..  62  Figure 5.2.3: MFCS-8 high precision pressure regulator……………………………..  64  Figure 5.2.4: Experimental setup during cell-loading process.………………….……  65  Figure 5.2.5: Cell viability comparison static condition vs. perfusion condition.…...  67  Figure 5.2.6: Syringe pump was used to consciously supply culture media....……….  68  Figure 5.3.1: HFL1 cell viability comparison…………...……………………………  72  Figure 5.3.2: HFL1 cellular shape……………………………..…………….………..  73  Figure 5.3.3: Hep G2 cellular shape…………………….…………………………….  75  Figure 5.3.4: The Hep G2 cell line inside the microchannel form 3-D spheroids..…..  76  Figure 5.3.5: Hep G2 cell viability comparison….……………………………….…..  77  Figure 5.3.6: LCC6/HER2 cellular shape…...……………..…………………………  78  Figure 5.3.7: The LCC6/HER2 cells form 3-D spheroids in the gel layer……………  79  Figure 5.3.8: LCC6/HER2 cancer cell viability comparison……..…………………..  80  Figure 5.3.9: The trapping process for co-culture two cell types simultaneously……  81  Figure 5.3.10: Co-culture HFL1 and Hep G2 cells…………………………….……..  82  x  Figure 5.3.11: Cell releasing…..……………………………………………………...  83  Figure 5.3.12: Cell releasing viability comparison……….…………………………..  84  Figure 5.3.13: The molecular structure of doxorubicin (Dox)… ……………….…....  86  Figure 5.3.14: Structures of Rhodamine B (C28H31ClN2O3)………………………….  87  Figure 5.3.15: Diffusion time measurement of Rhodamine B through alginate……...  88  Figure 5.3.16: Diffusion distance of Rhodamine B through PDMS……..…………...  90  Figure 5.3.17: Control experiment before drug testing ………………….…………...  92  Figure 5.3.18: Control experiment result…………………………………..…………  93  Figure 5.3.19: Cell viability reading for Dox drug testing…….…….………………..  94  Figure 5.3.20: Drug testing result…………………………………………………….  95  xi  Acknowledgments I would like to thank Dr. Karen Cheung for her inspiration, encouragement, patience and support over the last two and a half years. I am very fortunate to have this opportunity to work with her. I would like to thank Dr. Mu Chiao, Dr. Eric Lagally, Dr. Jayachandran Kizhakkedathu and Dr. Boris Stoeber for sharing their lab equipments and being generous with their time and knowledge. I would also want to thank Dr. Helen Burt and John Jackson from UBC faculty of pharmaceutical science for their generous support. I am also grateful to my dear friends: Vahid Bazargan, Jonas Flueckiger and Madhuja Gupta. This project would not have been possible without their help. We had a wonderful time together here. Finally, I wish to express my genuine gratitude to my family.  xii  Dedication  To my family  John Chen, Jane Chen and Robert Chen  xiii  CHAPTER 1  Introduction 1.1 From Semiconductors to Biomedical Microdevices BioMEMS stands for biological microelectromechanical systems.  It can be  defined as microelectromechanical systems (MEMS) used for biomedical applications or life science. MEMS technology is originally based upon the photolithographic process used to make computer chips [1]. This technology enables production of low-cost, highfunctionality devices for everyday use, like printer cartridges for ink jet printing and miniaturized accelerometers for automotive airbags. BioMEMS promise the production of miniature, smart, and low-cost biomedical devices that could revolutionize clinical practice with improvements to surgical instruments, artificial organs, diagnostic tools, and drug delivery systems [2]. For cell biological studies, microdevices allow uniform nutrient and oxygen diffusion. They are the perfect tools to conduct cell-based experiments. BioMEMS techniques have great  1  impact on applications like tissue engineering, cell separation and culture devices, and high-throughput drug screening tool and cellular bio-detection [3]. By miniaturizing medical or biological devices, it is possible to work on small sample sizes and consider each cell as a unique identity. BioMEMS could bring the concept of Micro Total-Analysis System (µ-TAS) to reality. µ - TAS integrates devices to create a system to automate complex operations like cell sampling, culture, monitoring and detection.  1.2 Micro Total-Analysis System (µ-TAS) The concept of Total-chemical-analysis system (TAS) first appeared in the 1980’s. The goal was to simplify the analytical steps in chemistry and had samples flown from one step to the other without human intervention [1]. The concept of Micro TotalAnalysis System (µ-TAS) was first introduced by Manz et al. in 1990 to describe advantages in chromatography and electrophoretic separation [4].  In 1991, Manz  proposed the idea of using photolithographic techniques for producing microdevices for electro-osmotic flow separation. At the same time, Verpoorte microfabricated the first silicon microdevice for cell detection [2].  2  Figure 1.2.1: Micro Total-Analysis System can incorporate 3-D scaffolds for guiding cell growth, and transporting nutrients and reagents. Multiple functionalities for biochemical analysis (such as image-based analysis) can be integrated on a single device. [Courtesy: Jonas Flueckiger]  Most µ-TAS and Lab on Chip devices (LOC) device are based on the concept of microfluidics, which is the study of transport processes in microchannels. These are channels with a hydraulic diameter below 1 mm [2]. electrophoresis was one of first devices of this type.  Chip-based capillary  Similar to microelectronics,  microfluidic technology makes it possible to fabricate highly integrated devices, which incorporate multiple components (valves, mixers, pumps and filters) for performing multiple tasks without human intervention (Figure 1.2.1). Some of the most common applications for µ-TAS and LOC devices include genomics, proteomics, clinical diagnostics, drug discovery and biosensors [5].  3  1.3 Cell Trapping in Microfluidic Chip Immobilizing and manipulating cells are the fundamental cell-handling operations (separation, concentration and manipulation), which are required for almost all biological researches [6, 7]. Trapping cells in microfluidic chips poses many challenges because of the complex physical properties of cells. Various trapping mechanisms have been developed Some of the most commonly used methods are electro-osmosis, electrophoresis, dielectrophoresis, optical interference, acoustic standing waves, splitting laminar flows, mechanical obstacles and magnetic forces [6, 7] (Figure1.3,1).  Figure 1.3.1: Cell-immobilization methods  4  1.3.1 Chemical Trapping Channel surfaces are modified in topography or chemical composition, providing a discrete location for cell adhesion or protein attachment [7]. In recent years, surface modifications have been integrated with microlithography and microfluidics to achieve high precision cell trapping [8, 9]. In order to achieve an effective trapping surface, the coating material must be highly effective for all deposited cell types. It should also inhibit cell migration and overgrowth. However, there are some drawbacks of surface immobilization methods. They are only applicable to adherent cells, and once attached, the process is often irreversible [7].  1.3.2 Hydrodynamic Trapping This type of trapping uses variations of surface topography to separate particles from a flow. Mechanical obstacles or barriers are tailored to the size of the particles to be captured, so the obstacles facilitate passage for fluids only [7, 10].  Besides using  mechanical barriers, regions of low shear stress can also be used for retaining particles next to the moving fluid [11]. This method has an advantage over other trapping methods for rapid cell immobilization.  In addition, these devices are often simple and  inexpensive. The shortcomings of this type of device are that contact of the cells with a surface is not avoidable, which might lead to irreversible attachment. Also it is difficult to achieve a single particle deposition in hydrodynamic trapping devices. Array sites are often either empty or filled with aggregates [7].  5  1.3.3 Dielectrophoretic (DEP) Trapping Polarizable particles such as cells, viruses, proteins or DNA molecules can be trapped within low-viscosity, low-conductivity solutions using dielectrophoretic trapping [12]. Dielectrophoresis is a phenomenon that uses a non-uniform AC electric field to exert a force on polarized neutral particles. The permittivity and conductivity of the particles and the liquid medium, varies as a function of the frequency of the applied AC field. Depending on the differences between these two values, trapped particles can either experience a positive (move toward the higher field) or a negative (repelled from the high field) dielectrophoretic force [13, 14]. Electric cages can be built that enable a single cell to be trapped within the flowing medium [15, 16].  A drawback of  dielectrophoresis trapping is that cells are constantly exposed to strong AC fields, which causes Joule heating and sets up transmembrane voltage. Many conventional cell culture biomaterials such as collagen and alginate are too conductive and viscous to be used in this trapping method [7, 12].  1.3.4 Optical Trapping An optical trap exerts two forces on a particle. One is the scattering force that drives the particle away from the light source by radiation pressure. The second one is the gradient force that pulls the particle into a high-intensity region of the light beam [7, 17]. Laser and infrared are some of the common light sources that are used in this trapping method [17]. To use optical trapping, particles need to be transparent. In addition, the refractive index of the particle should be different from that of the surrounding medium at the trapping wavelength. However, since each trap requires a  6  fixed amount of optical power, the number of traps are limited by the available power of the light source [7]. Another shortcoming is that the trapped cell can get severely damaged due to overheating by the light sources [17, 18].  1.3.5 Acoustic Trapping With acoustic trapping, particles are subject to the mechanical force of a standing acoustic wave field that is generated by integrated ultrasonic transducers [7]. Acoustic force results from different densities and sound speeds between the particles and fluid. The magnitude of the force is linearly proportional to the particle volume and acoustic frequency [19]. This technique has been used for particle concentration, agglomeration and separation [7, 18, 19].  1.3.6 Magnetic Trapping Magnetic trapping is applicable to diamagnetic particles as long as their magnetic susceptibility is different from that of the suspension [7]. Different configurations of magnetic traps can be fabricated on silicon or glass substrates using photolithography techniques to capture single cells [20]. To achieve high trapping efficiency, the cells can be magnetically labelled and the solution can be modified with paramagnetic ions. However, antigen labelling is an invasive technique and it denaturizes the sample involved [21].  7  1.3.7 Hydrogel Trapping Cells immobilized in a polymer matrix are often regarded as being between “contact” and “contact free” immobilization. Hydrogels, in general, are very hydrophilic in nature. For example, agarose has a water content of more than 99 %. Gel trapping allows the liquid medium to have access to the whole cell surface whereas in chemical trapping or hydrodynamic trapping, cells often lean against a flat surface [7].  An  advantage of this type of cell entrapment is that it does not require a continuous electromagnetic field or light source which prevents overheating and mechanical stress. Cells can be trapped and then cultured for extensive periods of time [7].  1.4 Traditional Cell Culture and 3-D Cell Culture in Microchannel  1.4.1 Traditional Cell Culture Traditionally, petri dishes, culture flasks and well plates are used to maintain both non-adherent and adherent cell lines. For adherent cell lines, the cells are attached to the bottom of the culture flask and remain covered by a layer of cell culture media. For nonadherent cell lines, the cells are suspended within the culture medium. It is a complex method involving an array of supplies like petri dishes, pipettes, and conical tubes. Recently, there has been increased interest in 3-D cell culture, in which cells are embedded in supporting structures that mimic the extracellular matrix (ECM) [22]. 3-D culture is biochemically and physiologically more similar to in vivo tissue, but is  8  technically challenging in terms of maintaining factors such as nutrient and waste exchange.  1.4.2 3-D Cell Culture in Microchannel The ability to culture and reconstitute tissue structures and functions in vitro has a tremendous influence on clinical applications [23, 24]. Within mammalian tissues, cells are supported by a network-like structure, known as the Extra Cellular Matrix (ECM) that give tissues their mechanical strength and helps organize cell-cell communication. Led by cancer researchers, biologists are increasingly excited about 3-D cell cultures, where they have discovered patterns of gene expression and other biological activities that mimic in vivo conditions more closely and cannot be observed with traditional 2-D cell culture methods [22, 25]. Cell culture using microdevices has an advantage over traditional culture systems in terms of scale, efficiency and cost [25-28].  For instance, by using a microchip as a  culture flask, rapid and secure exchange of culture media and reagents are possible while making continuous microscopic observations.  In conventional experiments using a  culture flask or petri dish, a long diffusion time is needed for added reagents to reach each cell and localized rapid chemical stimulation is nearly impossible [25, 29]. However, it is possible to realize rapid chemical stimulation with a microfluidic system [30]. In an integrated microfluidic chip, biochemical experiments can be carried out automatically by culture cells with chemical sensors [27, 28, 31]. Microfluidic systems which integrate 3-D cell culture techniques could provide a microenvironment that resembles the physiological environment more closely than can be  9  achieved by a traditional 2-D culture system.  By carrying out experiments at the  microscale, precise experimental parameters can be applied efficiently on small cell numbers or even to individual cells. This process will be beneficial in the areas of stem cell research and drug discovery. Some of the work that has been done with culture cells inside microfluidic devices is listed in Appendix A.  1.4.3 Co-culture Multiple Cell Types Our body operates as a society or ecosystem whose individual members are cells. Cell-cell interactions are critical to the development of multicellular organisms [32, 33]. Culturing multiple cell types within close proximity of one another has long been used in biological research. For stem cell research, a layer of feeder cells is traditionally cocultured with the stem cells.  The feeder cells provide ECM and specific factors  necessary for the maintaining of stem cells in the undifferentiated state without losing pluripotency [34, 35].  Other examples like interactions between endothelial cells and  muscle cells affect the expression of regulatory factors such as fibrinolytic and coagulation factors and angiogenic factors [36].  In pathological studies, cell–cell  interactions can also implicate diseases like atherosclerosis in cardiovascular disease, denvervation atrophy in skeletal muscle and alcoholic cirrhosis in liver disease [9].  10  1.5 Motivation of this Work Conventional cell studies are conducted with large populations of cells; therefore measurement can only reflect average cell response values.  This can lead to  misinterpretation, since some biological mechanisms of a cell are in binary state (“on” or “off”) [7]. For example, the majority of existing evidence suggests that, within individual cells, a gene is either maximally expressed or is not expressed at all [37]. Because of cellular heterogeneity, at an intermediate state a cell population will contain one portion in the “on” state and another in the “off” state. The relative numbers of “on” and “off” states often appear as a gradual change in the response of each individual cell. It is necessary to conduct genetic, physiological, and biochemical cell studies on a single cell and with a sufficient number of cells to obtain statistically meaningful data [7]. Traditionally, cell culture has been done with culture flasks, well plates, or petri dishes, which are many orders of magnitude larger than the cellular level. The culture flask is often continuously stirred or refreshed with cell culture medium. This action disrupts the microenvironment and creates stress on the cell due to stirring and oxygen bubbles [25, 38]. Culturing cells within a hydrogel matrix in a microfluidic device allows important factors to accumulate locally forming a stable microenvironment which is closer to in vivo conditions. It has been found that cells grow significantly slower in microchannels than in culture flasks [39].  This phenomenon is desirable in drug  screening, where repeated measurement is required to detect cell alteration resulting from disease or the application of xenobiotics. One of the major constraints on the size of in vitro tissue engineering is that there is no built-in blood supply; maintaining sufficient circulation of media can be  11  challenging. Oxygen and other nutrients can only diffuse through short distances before they are consumed [40, 41]. Microfluidic networks can be used for constructing and simulating blood vessels which mimic large-scale physiological properties [40]. In terms of expense, the use of microfluidic devices to conduct biomedical research is more economical compared to traditional cell culture methods. First, only a small amount of reagent and biological sample are needed to perform the experiment because the volume of fluids within the microchannel is very small (usually several nanoliters).  This represents a significant cost reduction, especially when expensive  reagents are required. Another advantage associated with small sample volumes is that the delivery and removal of soluble biochemical molecules in the extracellular microenvironment can be precisely controlled, which is desirable for highly parallel applications, such as drug discovery, genomics and proteomics [42].  Second, the  fabrication techniques used to construct microfluidic devices are compatible with the current photolithography techniques used for microchips and are relatively inexpensive. Different cell types are often co-cultured in close proximity within the same cell culture environment to achieve in vivo-like cell-cell interactions. Cell-cell interactions are important in the physiology of many organ systems such as the vasculature system (smooth muscle cell and endothelium) and the liver (hepatocyte and sinusoidal endothelium) [9]. Besides these interactions, cell-cell interactions are also implicated in the pathophysiology of many diseases. Many of the cellular interactions still remain a mystery [43]. One of the limitations of current co-culture systems is the inability to vary local cell-seeding density independently from the cell number [9]. By incorporating coculture with cell trapping in micro-fluid chip approach, the accuracy of manipulating cell-  12  cell interaction can be improved by precisely controlling cell placement inside the cell culture environment. Unlike other trapping methods such as laser trapping, acoustic trapping, magnetic trapping, and chemical trapping, immobilizing cells in hydrogels does not cause continuous shear stress on cell membranes and gives accurate control over the 3-D extracellular environment. This function can be further expanded to provide in vitro cellular co-culture and to better facilitate drug testing.  13  CHAPTER 2  2 Alginate Hydrogel Cell Encapsulation 2.1 Applications of Hydrogel Hydrogels are polysaccharides characterized by high water content and porous polymer networks. They have been used extensively in various applications because of their changing physical properties after exposure to different solvents, ions, pH levels, electric fields, temperatures, or light intensities [44].  The effect of expansion or  shrinkage is often utilized to construct artificial muscles or micro-valves in microfluidic systems. Other systems utilize porosity and high flow resistance of the polymer networks for analytical applications such as dialysis, electrophoresis, substance enrichment, and sample separation.  Besides the applications mentioned above, hydrogels are also  commonly used for cell trapping and drug delivery [44, 45].  14  2.2 Cell Encapsulation and Cultivation The human body is comprised of approximately 100 trillion cells, with more than 260 phenotypes. These cells divide, differentiate and self-assemble into tissues and organs [46, 47]. There is always a great need for living cells to restore and maintain the functions of our body. A variety of 3-D cell culture matrices, or scaffolds, have been developed for tissue engineering applications to restore or maintain tissue and organ functions (Figure 2.2.1) [46]. Unlike traditional cultures on flat surfaces, the 3-D scaffolds not only support cell viability but also metabolic activity and in vivo phenotypes.  Figure 2.2.1: Different cellular structure formation in 2-D and 3-D scaffolds [48].  15  2.2.1 Naturally Derived Hydrogel Naturally derived hydrogels, such as collagen, agarose, and alginate, have frequently been used in tissue engineering because they are either components of or have properties similar to the natural ECM [49].  2.2.1.1 Collagen Collagen is an attractive material for biomedical applications since it is the main component of natural ECM and comprises 25 % of the total protein mass [49]. Collagen is long fibrous structural protein composed of three polypeptide chains, which wrap around one another. The mechanical property of collagen fibres and scaffolds can be modified by introducing various chemical crosslinkers, physical treatments, and by blending them with other polymers.  Collagen can be naturally degraded by metalloproteases,  specifically collagenase, and the presence of cells [50].  2.2.1.2 Agarose Agar gels are made of polysaccharides derived from seaweed or algae. Agar gels have been used on a daily basis as food thickeners and as a vegetarian gelatine substitute. They also have been commonly used in bacterial cell cultures and in electrophoresis for DNA separation [32]. One of the main components of agar gel is agarose, a linear polymer with a molecular weight of between 106 400 and 243 500 g/mol [51].  The hydrogel is a  copolymer consisting of (1,3)-linked β-D-galactose and (1,4)-linked (3-6)-anhydro-α-L-  16  galactose. Agarose is a thermally reversible gel that solidifies around 35 °C and melts around 70 – 85 °C. The temperature depends on the polymer’s molecular weight and concentration [52]. Although agarose has also been investigated as a cell encapsulation material for transplantation, its use in immunoisolation has not yet been demonstrated [53, 54].  2.2.1.3 Alginate (Alginic Acid) Alginates are polysaccharides derived from seaweed. Similar to agarose, alginate has been commonly used as food stabilizers and thickeners, and as drug-releasing materials. Alginate is an ionically cross-linked gel. It consists of two types of copolymer blocks: (1-4)-linked β-D-mannuronic acid (M units) and α-L-guluronic acid (G units) [55]. The G units can be cross-linked with divalent cations such as Ca2+ or Ba2+ ion. This is a reversible process. The gel can be dissolved by the addition of chelating agents such as Ethylenediaminetetracetate acid (EDTA). Alginic acid has been extensively used in cell encapsulation. For example, islet cell transplantation using alginate microspheres has been investigated as an approach for treatment of insulin-dependent diabetes and hepatocyte encapsulation has been investigated in the development of bioartificial livers [56, 57].  2.2.2 Synthetic Hydrogel Synthetic hydrogels have the advantage of having controllable and reproducible chemistry and properties. They can be reproduced with specific molecular weights,  17  blocks structures, and crosslinking modes. These conditions, in turn, determine the gel formation and degradation dynamics [49]. Pluronic and N-isopropylacrylamide (NiPAAm) are some of the commonly used synthetic hydrogels.  2.2.2.1 Poloxamer Poloxamer, poly(ethylene oxide)-poly(propylene oxide)-poly(ethylene oxide) (PEO-PPO-PEO), is a reversible tri-block copolymer. It is also known by the trade name Pluronic.  Pluronic has been added to mammalian cell culture medium in low  concentrations as a surfactant from 0.05-0.1%. The surfactant lowers the surface tension of the medium to prevent cell damage due to shear stress and to improves the efficiency of metabolic transport [58, 59]. The synthetic hydorgel self-assembles into micelles due to the hydrophobicity nature of the tri-block structures, a central hydrophobic PPO block sandwich using two hydrophilic PEO blocks. At higher temperatures, aqueous pluronic solutions undergo thermally activated reversible gel formation. The micelles form into a tightly packed arrangement. The gelling temperature varies with the polymer concentration and its molecular weight. For 15 wt% pluronic F127, the critical solution temperature is around 28 °C [60, 61]. Pluronic F127 has been approved by the Food and Drug Administration (FDA) for drug delivery applications. It has also been used as a tissue-engineering scaffold material for chondrocytes, epithelial cells, and fibroblasts [62, 63]. For better mechanical and chemical stability, pluronic can be modified to add covalent crosslinking [64].  18  2.2.2.2 N-isopropylacrylamide Polymers (NiPAAM) N-isopropylacrylamide (NiPAAm) is a temperature-responsive polymer also known as PNIPA, PNIPAAM, PNIPAA or PNIPAm [65]. Similar to pluronic, NiPAAM is also a thermosensitive gel that exhibits a reversible phase transition [66]. In aqueous solutions, NiPAAM has a lower critical solution temperature (LCST) of 32–34 °C. When below the LCST, NiPAAM chains are hydrated and adopt a flexible and extended coillike structure. Above the LCST, NiPAAM chains become dehydrated and collapse into a tightly packed conformation [67]. NiPAAm has already been used for 3-D culture of chondrocytes and pancreatic islets, and as a carrier for controlled drug delivery [63, 66, 68-70].  2.3 Chemical Structure of Alginate Hydrogel Alginates (algins, alginic acids) are polysaccharides extracted from brown seaweeds such as Laminaria hyperborea and Lessonia. The first characterization of alginic acid was done by Stanford (patent 1881) [55].  It consists of two kinds of  monomer units: (1,4)-linked β-D-mannuronic acid (M) and (1,4)-linked α-L-guluronic acid (G) (Figure 2.3.1) [55]. These two kinds of monomers are covalently linked in various arrangements along the polymer chain (e.g. -M-M-M-M-, -G-G-G-G- or -G-GM-M-) (Figure 2.3.2). Alginate is typically composed of 100-3000 building units linked together with molecular weights of between 50 - 100,000 kDa [55].  19  b)  a)  COO  - OOC O  -  O  O  OH  HO HO  O  O  O  HO  Figure 2.3.1: Monomers of alginic acid: a) (1, 4)-linked β-D-mannuronic acid (M) and b) (1, 4)-linked α-L-guluronic acid (G).  - OOC O HO  - OOC  a)  O  O  HO  HO  HO  O  HO  O - OOC  HO  O  O - OOC  -  COO  HO  O  HO  HO  -  COO  HO O  O  O  OH  OH  b)  O OH  O OH  COO  COO  O  OH  COO  -  -  HO  OH  - OOC  O  O O OH  OH  -  O  O  c)  O  O  O  O  O  HO  HO  O  HO O - OOC  OH  COO  O  HO  -  Figure 2.3.2: Molecular structure of alginic acid: a) -M-M-M-M-, poly M segment. b) -G-G-G-G-, poly G segment. c) –G-G-M-M-, mixed segment.  20  The arrangement of these two components and the relative amount (G/M ratio) depends on the source from which the alginate was derived.  Figure 2.3.3: Bonding sites of Ca2+ ion. Only G blocks provide reactive carboxylic bonding sites for the Ca2+ ion. The Ca2+ ion attaches two different alginate strands together to form solid gel matrixes.  Two G blocks of adjacent polymer chains can be cross-linked with divalent cations, such as Ca2+ or Ba2+, through interactions with the carboxylic groups in the polymer chain to form a gel network (Figure 2.3.3). Thus, overall gel stiffness and pole size are dependent on the polymer G/M ratio. High M content alginate chains are more  21  flexible than high G content alginate chains [4, 55]. The diffusivity within the hydrogel is lower in the high G alginate than in the low G alginate due to a smaller pole size [71]. The reaction between sodium alginate and calcium should be expressed as follows:  The reaction is very rapid. Calcium alginate forms on the surface almost instantly when the sodium alginate solution comes in contact with the Ca2+ ion.  2.4 Mechanical Properties of LF120M Sodium Alginate The alginate used for this experiment was the LF120M type sodium alginate from FMC Biopolymer, Norway. The G/M ratio is around 35-45/55-65 %. With 1.0 wt% solution, the viscosity is around 150 mPa·s at 20 °C [72]. Alginate solutions are shear-thinning, non-Newtonian fluids. decreases with increasing shear rate.  The viscosity  The viscosity is dependent on solution  concentration, polymer chain length, and solution temperature. At similar concentrations, the shorter the polymer chains the lower the viscosity. The viscosity also decreases with increases in temperature. For mammalian cell culture, all labwares or materials used need to be sterilized to prevent infection and contamination. One of the most common methods of sterilization is autoclaving. Following the biology protocol, all solution used for the experiment, which includes the LF120M alginate solution, was heated to 121 °C  for 20 minutes to  inactivate all bacteria, viruses, fungi, and spores [73]. Alginic acid is a covalently-linked  22  long chain polymer. By autocalving, the polymer chain will be broken and the viscosity will be lowered. The polymer chain analysis showed that before and after autoclaving, the average molecular weight (Mn) of LF120M changed from 140,500 to 83,470 g/mole (measured with Gel Permeation Chromatography). The measured result was consistent with the viscosity measurement shown in Figure 2.4.1.  Figure 2.4.1: The viscosity of 0.5 wt%, 1.0 wt% 1.5 wt% and 2.0 wt% LF120M alginate solutions from 20 °C to 60 °C was measured with a Bohlin C-VOR rheometer. Each alginate solution was prepared in two different conditions: unautoclaved and autoclaved at 17 psi and 120 °C for 20 minutes. After autoclaving, the solution was noticeably less viscous.  23  2.5 Alginate Gel Stability It has been found that gel strength may vary with the kind of divalent cation used in the following order: Pb2+ > Cu2+ = Ba2+ > Sr2+ > Cd2+ > Ca2+ > Zn2+ > Co2+ > Ni2+ [74]. However, in applications involving immobilization of living cells, toxicity is a limiting factor in the use of most ions, and only Sr2+, Ba2+ and Ca2+ are considered nontoxic for these purposes. A long-term alginate gel stability test was carried out. In this test, 1.5 wt% alginate solution was bought in contact with 40 mM and 100 mM of the CaCl2 solution. Gel degradation could be affected by changes in polymer molecular weight (MW) [75], which can be altered during the sterilization process. Therefore, two of the most common sterilization protocols, autoclaving and UV sterilization, were incorporated as part of the test condition. A 200 µm wide gel slab was first formed inside the microfluidic channel and the channel was filled with a F12K cell culture medium. The microfluidic chip was placed inside the incubator at 37 °C and 5 % CO2 level. The cell medium inside the channel was changed three times a week. The experiment showed that the LF120M gel type is very stable under these cell culture conditions.  There was no sign of gel  dissolution throughout the two-week period from visual measurements (Figure 2.5.1). However, the mechanical strength of alginate gel is expected to degrade over time [53].  24  Figure 2.5.1: Stability of alginate in microchannel. The change in alginate gel layer thickness was measured every second day for 11 days. Alginate solutions were pretreated with two different sterilization protocols: a) autoclaved at 121 °C for 20 minutes or b) exposed to UV light for 2 hours. 100 mM and 40 mM CaCl2 solutions were used for each test condition. We found no signs of gel dissolution (0% change in gel thickness) from visual measurements.  25  CHAPTER 3  3 Microfabrication Technology 3.1 Polydimethlsiloxane (PDMS) Microfluidic Chip In recent years, microfluidic systems have attracted a lot of scientific and industry attention. Early development is mainly based on conventional semiconductor materials and fabrication technology intended for integrated circuits. Using these technologies and materials is costly and imposes many limitations on fabrication and testing. Several alternative technologies using organic polymers have been investigated, one of which is polydimethylsiloxane (PDMS) [76-78]. Besides the advantages of being biocompatible, permeable to gas, and inexpensive, PDMS is also transparent to visible and UV light wavelengths ranging from 230 nm to 700 nm. This is ideal for biological applications since most of the fluorescence markers and the UV lights used for sterilization (~ 265 nm) are well within that range [77, 79]. In addition to these, PDMS has a low curing temperature (<100 °C), which can be easily achieved by conventional oven. 26  In combination with photolithography technology, 3-D PDMS microchannels can be made easily and rapidly by the replica moulding method. This technique is costeffective, fast, and simple. A microchannel design can be turned into an actual device within 24 hours. Typically, 3-D channels are formed by exposing both PDMS layers to oxygen plasma and then bonding them immediately to a flat surface after the plasma treatment (see section 3.2) [78].  3.1.1 Chemical Structure and Property of PDMS PDMS is one of the most commonly used silicon-based organic polymers. The chemical formula of PDMS is (H3C)3SiO[Si(CH3)2O]nSi(CH3)3, where n is the number of the repeating monomer [SiO(CH3)2] [80].  The structure of PDMS is shown in  Figure 3.1.1.  Figure 3.1.1: The backbone structure of PDMS  3.1.2 PDMS Curing PDMS is cured by an organometallic crosslinking reaction between a base and a crosslinking agent [81]. A 10:1 weight ratio of base and crosslinking agent is typically used [82, 83]. The base, with a length of about 60 monomers, consists of a linear oligomer containing vinyl groups (-HC=CH2) at the terminations [84]. The crosslinking  27  agents, with about 10 building blocks, sometimes have a hydrogen atom attached to the Si atom instead of a methyl group. This bonding site is designated as R in Figure 3.1.2. The curing agent also contains a platinum-based catalyst that catalyzes the addition of the Si-H bond across the vinyl groups, forming Si-CH2-CH2-Si linkages [81, 84].  Figure 3.1.2: PDMS curing. The crosslinking agent consists of a siloxane oligomer with a length of about 10 units. At some of the bonding sites (R), the silicon atom has a hydrogen atom attached to it instead of a methyl group. The hydrogen atom will then react with the vinyl group of the base.  The properties of the cured PDMS are dependent on the ratio between the base and curing agent. The higher the ratio of the curing agent added, the harder the cured  28  PDMS becomes because more cross-linked elastomer is formed. After curing, the top and bottom of the PDMS layer remain smooth and flat. For proper curing, the base and crosslinking agent need to be thoroughly mixed. The weighted PDMS is mixed with an AR-250 mixer from Thinky for five minutes and cured over a master in an aluminum foil dish. Dow Corning Sylgard 184 PDMS can be cured over a wide range of times and temperatures, varying from ~48 hours at room temperature to 10 minutes at 150 ºC [85]. The crosslinking time can be shortened with higher curing temperatures.  3.1.3 Diffusivity of PDMS Chip  3.1.3.1 Oxygen Supply Oxygen supply is one of the most important parameters for cell culture. Because PDMS is highly porous, and microbioreactors have a high surface area to volume (SAV) ratio, the oxygen supply to the cultured cells from the atmosphere is sufficient; hence, there is no need to use an oxygenated culture medium [86]. The oxygen supplied by diffusion through the PDMS layer can be estimated with Fick’s first law: J = − DPDMS  ∆CO2 ∆x  (3.1)  where ∆x , ∆CO2 and D PDMS denote the thickness of the PDMS chip, the oxygen concentration difference across the PDMS layer, and diffusivity of oxygen through the PDMS layer. D PDMS is estimated as 4.1x10-5 cm2/s [87, 88] and ∆x equals 4 mm as measured. The typical oxygen concentration in a tissue culture medium is 0.22 µmol/ml,  29  assuming all oxygen is consumed by the cultured cells inside the channel so ∆C o 2 equals 0.22 µmol/ml [86, 89].  Based on these parameters, J is estimated to be  2.25 × 10-5 µmol/cm2·s. The fabricated channel has a surface area of 0.1 cm2. The amount of oxygen diffusing through this PDMS microfluidic device is calculated to be  1.353× 10-4 µmol/min. The oxygen consumption rate of MCF-7 human breast tumor cell type (6 nmol/min per 1million cells) [90] is used as an estimate for LCC6/HER2 breast tumor cell type used in this experiment. Since the initial number of cells cultured in this microfluidic device is from 500 to 1000 cells (section 5.2.1), the estimated oxygen supply is more than an order higher than the oxygen consumption by the cultured cell lines. It can be assumed that a sufficient amount of oxygen can be supplied to this device by simple diffusion at the initial stage. As the cell proliferates, the culture medium needs to be circulated through the culture chamber to ensure sufficient oxygen is provided to the cells.  3.1.3.2 Nutrient Supply Besides oxygen, the second most important nutrient in the cell culture is glucose. It provides the source of energy for metabolism. Even though PDMS is highly oxygen permeable, the same cannot be said about glucose. The permeability of PDMS (a highly hydrophobic polymer) to glucose is very low [91, 92]. Given that glucose is water soluble, one of the possible explanations is due to the hydrophobic nature of PDMS. The permeability of water into PDMS is generally considered very low [93]. Based on this  30  finding, the circulation of culture media through the microchip is essential for long-term cell culture. Nutrient supply through continuous perfusion is discussed and demonstrated in section 5.2.3.  3.2 PDMS Bonding The PDMS moulding process provides three of the four walls necessary for forming an enclosed channel as shown in section 4.4.2.2. The fourth wall is provided by bonding the PDMS replica to a flat surface. Bonding of a PDMS-based device is simpler than what is required for glass or silicon because high temperatures, pressures, and voltages are not required [94]. The adhering process can be reversible or irreversible [76].  3.2.1 Reversible PDMS Bonding Reversible bonding occurs when a flexible PDMS piece is in contact with a flat surface making a weak van der Waals attraction [76, 95]. This method is fast and occurs at room temperature. The sealing is watertight but cannot withstand high fluid pressure (< 5 psi) [95].  3.2.2 Irreversible PDMS Bonding One of the most common techniques for irreversible PDMS bonding to another flat surface is by oxygen plasma treatment. The flat surface can be either PDMS, glass, silicon, quartz, polystyrene, polyethylene or silicon nitride [76, 81].  31  PDMS contains repeating units of [SiO(CH3)2]. The methyl groups, –CH3 at the PDMS surface are turned into silanol groups, –OH, when exposed to oxygen plasma. Since the silanol groups are polar in nature, they will make the treated PDMS surface temporarily hydrophilic. When these silanol groups are brought in contact with those on another surface, they condense and form a covalent O–Si–O bond after the loss of water molecules (Figure 3.2.1). This covalent bond is irreversible [76, 81, 95, 96].  Figure 3.2.1: PDMS-PDMS bonding. After exposure to oxygen plasma, the methyl groups –CH3 at the PDMS surface are turned into silanol groups –OH. When these silanol groups are in contact with one another, they condense and form a covalent O–Si–O bond after the loss of water molecules. A similar process can be applied between a PDMS surface and another flat surface made of either glass, silicon, quartz, polystyrene, polyethylene or silicon nitride [81].  The bonding strength is affected by the RIE power and exposure time during plasma treatment. At low power levels, the kinetic energy of ions incident on the substrate reduces. Low power level also reduces the electron acceleration within the plasma environment leading to a reduction in the radical density. As a result, less active sites (Si-OH groups) are available for surface bondage. With higher power levels, there  32  is an increase in the ion bombardment thus damaging the siloxane backbone. Overexposure may also cause an increase in silanol bond density and surface chain scission reactions, resulting in surface reorientation and cracking [95-97]. The damage is unrecoverable. The same can also be said about exposure time. After the oxygen plasma treatment, PDMS becomes hydrophilic. hydrophobicity recovers if the treated samples are left in the air.  However,  The recovery  mechanism is not yet fully understood. Some of the possible reasons are migration of a low molar mass species from the bulk to the surface, reorientation of polar groups at the surface into the bulk, condensation of silanol groups at the surface, external contamination of the treated surface, changes in surface roughness, and loss of volatile oxygen-rich species to the atmosphere [81, 96].  The surface wettability can be  considered one of the indicators of bonding strength. A contact angle below 5 degrees is a general requirement for good bonding [95]. This implies that the bonding process should be carried out as soon as possible.  3.3 SU-8 Photolithography Polymer micromolding is a common way to make microchannels because of the ease of processing, low cost, and bio-compatibility [98]. The moulds often require sub micron resolution and high-aspect-ratio.  EPON SU-8, an epoxy-based negative  photoresist, originally developed by IBM and sold by Shell Chemical, is ideal for this application [99, 100]. It has low molecular weight and can be spin-coated as very thick films up to 500 µm in a single coat. The photoresist can also assemble structures with thickness of up to 2 mm [81]. Similar to other types of negative photoresists, SU-8 has  33  good adhesion to various substrates such as Si, GaAs, InP, and glass. Additionally, it exhibits excellent thermal and chemical stability.  Figure 3.3.1 shows the chemical  structure of SU-8.  Figure 3.3.1: Molecular structure of SU-8. SU-8 is a highly functionalized molecule with 8 epoxy groups. After exposure to ultraviolet light, e-beam or x-ray, the epoxy resist will form a highly structured cross-linked matrix [101].  SU-8 belongs to the category of chemically amplified photoresists. Chemical amplification increases the sensitivity to the exposure energy.  In the process, acid  released by the exposure radiation diffuses during the post exposure bake step. The acid renders surrounding polymer soluble in developer [102, 103]. A single acid molecule can catalyze many deprotection reactions; hence, fewer photons or electrons are needed [104]. Acid diffusion is important not only to increase photoresist sensitivity, but also to ease edge roughness.  34  The absorption of SU-8 in the UV spectrum is very low, thus allowing the epoxy resin to achieve high aspect ratio; aspect ratio > 20 by using UV light in the 365 to 436 nm range, and >80 by using electrons and x-rays. Exposure introduces a high level of cross-linking density, converting the SU-8 photoepoxy into a strong polymer [99, 100, 103].  SU-8 can be dissolved by solvents such as most ketones, toluene, and diacetone  alcohol. The SU-8 patterning procedures are listed below [81, 100, 103, 105]: 1.  Wafer Preparation: The wafers are cleaned with a RCA-1 cleaning procedure. Si  wafers are placed in the Piranha solution, a 3:1 mixture of sulphuric acid (H2SO4) and hydrogen peroxide (H2O2). When finished, the wafers are thoroughly rinsed with DI water. The cleaned Si wafers go through a dehydration bake to eliminate any moisture trapped inside the substrate. The Si substrate can also be cleaned with oxygen plasma. This treatment would also increase the adhesion of the photoresist to the wafer. 2.  Spin coating SU-8 photoresist: To have a uniform layer of epoxy, the SU-8  photoresist is spun for at least 40 sec. Because SU-8 2075 from MicroChem is highly viscouse, the wafer must go through a ramping step before reaching the final spin speed. The thickness of the coating depends on the spin speed and viscosity of the epoxy. 3.  Soft bake:  The soft bake evaporates the solvent and cures the epoxy before  exposure. This prevents the substrate from sticking to the mask. The baking time varies according to the thickness of the feature. Mild temperature ramping is required to prevent bubbling of the resist film.  35  4.  Exposure:  UV radiation is used to initiate the cross-linking of the SU-8  photoresist.  The exposure time has to be adjusted depending on the coating  thickness. The exposure pattern is defined by the mask, which defines which areas of the resist surface will be exposed to radiation and those that will be covered. 5.  Post Exposure Bake (PEB):  SU-8 is belongs to the category of chemically  amplified photoresist. The PEB catalytically completes the photoreaction initiated during exposure and reduces mechanical stress formed during soft bake and exposure. 6.  Development: Development involves the washing away of the non-polymerized  SU-8. The developer solution is made of PGMEA (propylene glycol methyl ether acetate). The development time is dependent on the thickness of the feature. The developed wafer is washed with isopropanol and blow-dried with nitrogen gas. 7.  Hard Bake: Hard bake may be applied after development to improve thermal,  physical, and chemical stability.  Because of the different thermal expansion  coefficient between resist and substrate, gentle temperature ramping and slow cooling are required to prevent cracking.  36  CHAPTER 4  4 Microfluidic Chip Design 4.1 Drag and Laminar Flows A PDMS-moulded channel is often rectangular in shape. In order to approximate the flow through the rectangular channel (where the height and width are comparable), the concept of hydraulic diameter Dh is used. The hydrodynamic diameter is given by: Dh =  4× A Pwet  (4.1)  Hydrodynamic diameter allows engineers to approximate flows though irregular channel shapes as those flows through channel with circular cross section. Pwet stands for wetted perimeter, which is the perimeter of the channel that is in direct contact with the flow, while A is the cross-section area where the flow goes through.  If the channel is  completely filled, Pwet and A are simply the diameter and cross-section area of the channel [106].  37  For flows in a long straight channel with a constant cross-section, Reynolds number, the ratio of the inertial forces to the viscous forces, is a good indication of whether the flow is laminar or turbulent [106-108]. The Reynolds number is given by: Re =  ρDhu Dhu = η v  (4.2)  As discussed above, Dh is the hydrodynamic diameter. u is the mean flow velocity of the flow in the channel, η is the dynamic viscosity of the fluid, v is the kinematic viscosity, and ρ is the density of the fluid. Table 4.1-1 lists the relationship between the Reynolds number and flow profile for circular pipe flow.  Table 4.1-1 Reynolds Number. It can be used to determine if flow is laminar, transient or turbulent. The information is for circular pipe flow. Re < 2000  Laminar Transition (Turbulent or  2000 < Re < 4000 Laminar) Re > 4000  Turbulent  For our application, most of the solutions used have properties similar to water (dynamic viscosity = 0.001 Pa·s, density = 1,000 kg/m3). The designed microchannel was 90 µm high, 800 um wide and 1000 um long, which gives the Dh of 161.80 µm. The mean flow velocity of the injected solutions was controlled at around 200 µm/sec.  38  With this information, the Reynolds number can be calculated ( Re =0.03236 ≈ 10-2). With a Reynolds number <<1, the flow profile remains laminar as shown in Figure 4.1.1.  Figure 4.1.1: Laminar flow in the microfluidic chip. By applying appropriate settings (mean flow velocity ≈ 200 um/sec and solutions with dynamic viscosity similar to water), we can create multiple streams of laminar flow inside the microfluidic device.  4.2 Cell Trapping By using the phenomenon that an alginate solution will form a 3-D gel matrix after contact with the Ca2+ ion, we can trap various cell types inside. The microfluidic chip contained a minimum of three inputs: 40 mM CaCl2 solution, 40mM EDTA solution and 1.0 wt% alginate solution.  39  The input ports were connected to a pneumatic flow control unit (see section 5.2.2).  Because of a much higher viscosity of 1.0 wt% alginate solution  compared to the CaCl2 solution at room temperature, CaCl2 was injected at a higher flow rate than alginate to keep the interface near the center of the channel.  d)  a)  b)  c)  Figure 4.2.1: Operation principle. a) Cells were suspended in alginic acid (1.0 wt% LF120M buffered in tris-HCl). We used the other two inlets for EDTA and CaCl2. At the loading step, a laminar flow of 40 mM EDTA was used to separate the Ca2+ ion from contacting with the alginate flow. b) After the EDTA flow was suppressed, the 40 mM Ca2+ ion solution was brought in contact with the alginate phase. The gel matrix started to grow from the boundary into the alginate stream with cells trapped inside. c) Once the gel formation step was completed, cell medium was introduced into the channel. d) The actual picture taken from the microfluidic device with 3 µm fluorescence beads added to the alginate solution.  40  The cell suspension was prepared with 1.0 wt% LF120M alginate solution and culture media. At the initialization stage, EDTA was used as a separation fluid to prevent unwanted gel formation and channel clogging. Once the steady state was reached, EDTA flow was suppressed and the alginate solution was brought in contact with the Ca2+ ion. The gel matrix started to form at the boundary as the gel slab grew toward the alginate flow stream with the cells trapped inside. All solutions, except culture media, were prepared with 50mM tris-HCl buffer solution with a 7.8 pH level and passed through a 5.0 µm Millex syringe filter (Ireland) to remove particulates during preparation before autoclaving. After the cells were trapped, the culture medium was introduced into the channel for long-term cell culture. The whole cell-trapping process is shown in Figure 4.2.1.  4.3 Diffusion and Gel Growth  4.3.1 Gel Formation Depending on the degree of polymerization, the average molecule weight of alginate is in the rage of 50 - 100,000 kDa, and the molecule weight of the Ca2+ ion is 40.08 Da. Since the molecule weight of the alginate polymer is much heavier than the Ca2+ ion, the gel formation was controlled by the speed in which the Ca2+ ion diffuses from the contact interface through the alginate gel block and reacts with the alginate to form new gel layers (Figure 4.3.1).  41  a)  b)  c)  d)  Figure 4.3.1: Gel formation a) 10 seconds, b) 30 seconds, c) 50 seconds, and d) 70 seconds after alginate phase was brought in contact with CaCl2. The picture shown is done with 1.0 wt% LF120M alginate gel with 40 mM CaCl2 solution. 0.7 µm fluorescent beads were added to the alginate solution as an indicator.  The relation between the formed gel thickness and time is shown in Figure 4.3.2, where the x-axis represents the time in seconds and y-axis represents the formed gel layer thickness in micrometers. The result of the graph can be explained with Fick’s first Law:  J = −D  ∆C ∆x  (4.3)  where J is the mass flux and ∆x represents the distance that the Ca2+ ion is diffusing through. D is the diffusion coefficient of Ca2+ ion in the newly formed gel layer and ∆C is the concentration gradient between the calcium stream and alginate stream. Since  42  we assume all the Ca2+ ions are consumed during gel formation (zero concentration at gel-alginate boundary), ∆C will equal the calcium solution concentration (cbulk(CaCl2)). If we assume gel growth rate is slow compared to diffusion times of ions in aqueous solution and we have approximately a linear concentration gradient of free Ca2+ ions (plus the ones bounded to alginate, but they don’t count for the diffusion calculation) across the gel layer, the quantity of Ca2+ ions that diffuses is:  cbulk (CaCl2 ) dnCa 2 + =D ×A ∆x dt  (4.4)  A is the cross section area where the calcium ions diffuse through. The quantity of Ca2+ needed to make a new layer of gel depends on the thickness of the new layer:  dnCa 2 + = dx × A × q  (4.5)  q is the volume density of Ca2+ binding sites. By rearranging equation (4.4) and (4.5), and combining all constants into p. Following expression for gel growth rate can be derived:  dx p = dt x  (4.6)  The constant p takes the form of diffusion constant. A higher ion concentration (cbulk(CaCl2)) means a larger p value, which results in faster gel formation. Equation (4.6) can only be considered as an approximation. For a more accurate model, the diffusion time of calcium ion in alginate gel and the effect of convection should not be ignored ( D ≈ 7.92 × 10−10 m 2 / s at 25°C in aqueous solution [109]).  43  Figure 4.3.2: Alginate gel growth rate increased with higher Ca2+ ion concentrations. The 1.0 wt% LF120M solution was prepared with 50 mM tris-HCl pH 7.8 buffer solution. The growth rate decreased with increasing gel thickness.  4.3.2 Gel Dissolving The goal of this project was to design a microfluidic device that can perform reversible cell trapping. After cells are trapped and cultured inside the alginate gel, they can be released by reintroducing EDTA back into the channel.  EDTA is a strong  chelating agent which binds to metals ions (Ca2+) via four carboxylate and two amine groups. By reintroducing EDTA, the chelating agent will deprive the calcium ion and  44  dissolve the alginate gel. Unlike the gel formation, where the calcium ions diffused through the gel block and formed new gel layers at the opposite flow stream, EDTA dissolved the gel block at the EDTA / gel slab interface. If we maintained a continuous flow stream of EDTA solution, the gel block was dissolved at a constant rate. As the EDTA concentration increased, the dissolving rate increased (Figure 4.3.3). The dissolving rate can be fitted in a straight line:  dx = −r dt  (4.7)  dx / dt represents the change of gel thickness over time and r is the gel dissolving rate.  Figure 4.3.3: Alginate gel dissolving rate. Alginate gel can be dissolved by reintroducing EDTA, a chelating agent, back into the channel. A 160 µm gel layer was preformed within the channel for this test. The experiment was done using various concentrations of EDTA solution, which were buffered with tris-HCl. The average flow speed of EDTA stream was ≈ 150 µm/s.  45  4.4 Microfluidic Chip Design Realization  4.4.1 Chip and Mask Design There are two main considerations for designing the fluidic chip: main channel dimension and the number of inlets. The device was designed for trapping and long-term culture of mammalian cell lines, LCC6/HER2 breast cancer cells, HMVEC endothelials, HepG2 hepatocytes, and HFL1 fibroblasts. The cell size varied from 10µm to 40µm in diameter.  Since LCC6/HER2 and HepG2 cell lines proliferate and form 3-D clumps  inside the gel matrix (section 5.3), the channel height was designed be at least 3 to 4 times the size of the used cell line to prevent clogging, minimize shear stress and provide enough cell culture space. The width of the channel was intended to fit the microscope viewing window and prevent channel collapsing. There were two types of channel patterns used. One had three independent inputs and the other one had five independent inputs. The patterns are shown in the figure below (Figure 4.4.1).  46  b)  a)  Figure 4.4.1: Channel design: a) 3 input and b) 5 input design used in this project. The main section of the channel was 800 µm wide, 90 µm high and 1 cm long.  The 3-inlet design was mainly used for gel characterization such as gel forming and dissolving rate measurement. The 5-inlet design was used for single cell type or multiple cell type co-cultures.  4.4.2 Micro-Fabrication Process  4.4.2.1 SU-8 Master Mould The designed microfluidic device with minimum features of 100 µm,was fabricated in the AMPEL nanofabrication facility, University of British Columbia (UBC). Because SU-8 is highly transparent to i-lines (365 nm), the exact wavelength provided by the cleanroom mask aligner, the patterned sidewall can be very vertical for thicknesses of up to 100 µm.  The epoxy also exhibits excellent chemical stability,  adhesive strength and resistance against stress corrosion [110]. It is the ideal patterning material to use as the master mould for our PDMS chip.  47  For our fabrication process, the following cleanroom equipment and material was used:  •  Canon PLA-501F double-side 100 mm mask aligner: It is designed to work with  4” wafers. The light source was a Hg lamp (USHIO 250W, Super High Pressure Mercury Lamp) with the wavelength at i-lines (365 nm).  •  Laurell WS-400B Spin Processor: The spinner provides automated ramping  functions with a maximum spin speed of 8000rpm. It is ideal for photo-resist coating.  •  Trion Plasma Enhanced Chemical Vapour Deposition (PECVD): The machine  comes with a 300 watt, 300 kHz bottom powered electrode. It was used for the wafer cleaning process and PDMS bonding with a glass substrate. For mould fabrication, we used the standard SU-8 fabrication process mentioned in section 3.3. SU-8 2075 from MicroChem, USA was chosen because of its ideal operating characteristics for a feature height of around 100 µm. The recipe used for fabrication is shown in Appendix B.  4.4.2.2 PDMS Channel The PDMS used was the Sylgard 184 from Dow Corning.  The fabrication  process followed the procedure discussed in section 3.1. The base and crosslinking agent (10:1 ratio) were thoroughly mixed before pouring onto the master. Before placement inside the oven, the samples were placed inside desiccators for removing air bubbles created during pouring. Dow Corning Sylgard 184 PDMS can be cured over a wide range of temperatures as discussed in section 3.1.2. For our experiment, the PDMS was placed inside the oven  48  at 50 ºC for 24 hours. The curing temperature was set at a lower temperature to prevent the SU-8 pattern from cracking due to the different thermal expansion coefficient between the SU-8 pattern and Si wafer. Based on experience, the PDMS was cured within 2 hours at 50 ºC. Applying a longer baking time was to ensure thorough curing and prevent uncured PDMS chain effects biocompatibility (section 5.1.2). For PDMS-glass bonding, the Trion PECVD machine located in the UBC cleanroom was used. The microchannel fabrication process is shown in Figure 4.4.2 and Appendix B The completed chip is shown in Figure 4.4.3. The finished chip was 1.5 cm wide, 2.5 cm long and 0.3 cm high, which fits perfectly into standard 6-well plates.  49  a)  e)  b)  f)  c)  g)  d)  h)  Figure 4.4.2: The fabrication process of the microfluidic chip: a) spin coating photoresist, b) UV exposure, c) post exposure bake, d) developing, e) PDMS curing, f) mould release, g) punch input and output holes, and h) PDMS-glass bonding.  50  Figure 4.4.3: The finished chip was 1.5 cm wide, 2.5 cm long and 0.3 cm high with a main microchannel that was 800 µm wide, 90 µm high and 1 cm long. The channel was injected with blue dye for visualization.  4.4.3 Polyethyleneimine Coating (PEI) Hydrogels typically have weak adhesion to surfaces; the same can be said about calcium alginate on glass substrate [44]. When gel adhesion is weak, bending and shifting of the gel slab is observed. This uncontrolled gel movement is undesirable during various stages of the experiment. The shifting of the gel wall causes a leakage flow of the calcium solution between the gel and channel wall, which results in an irregular gel shape during gel formation. Here, the culture media needs to be circulated through the culture chamber for long periods of time; a weak gel-surface linkage causes cell samples to be flushed out in the middle of the experiment. At the end of the research stage, alginate gel was dissolved and cells are released in a controlled manner. Instead of a gentle release, weak adhesion causes samples to be washed away in a sudden burst.  51  In order to prevent the problems mentioned above, improving adhesion between the alginate gel and glass surface was necessary. It can be done by coating the surfaces with a thin layer of polyethyleneimine (PEI). PEI is a cationic, non-toxic organic polymer, which is commonly used in DNA analysis and transfection of eukaryotic cells [111-113]. It is water soluble and has a strong affinity for protons due to a large number of nitrogen atoms of amino groups [114, 115]. The basic chain structure of PEI is [-CH2 -CH2 –NH-] n. In aqueous solutions, the nitrogen atoms of amino groups combine easily with H+ to form [-CH2 -CH2 -NH2+ -]n. The chemical structure is shown in Figure 4.4.4. Since alginate is negatively charged, there is a strong interaction between positively charged, PEI-coated surfaces and alginate gel layers [44].  Figure 4.4.4: Molecular structure of PEI. The basic chain structure of PEI is [-CH2 -CH2 –NH-] n. In aqueous solutions, the nitrogen atoms of amino groups combine with H+ to form [-CH2 -CH2 -NH2+ -]n.  52  For coating the sample, an aqueous solution of 0.5 wt% PEI (50 % (w/v), MW 750,000, Sigma Aldrich) and 1 M NaCl (the addition of salt into polyelectrolyte system causes deposition in thicker layers of polyelectrolyte chains [116]) was prepared and used to fill the channel for 24 hours following the bonding of the microfluidic chip. Afterward, the channels were thoroughly rinsed with water with a syringe pump at the rate of 0.01 ml/min for 20 min. The excess PEI solution was removed; otherwise, it would cause polymerization of the alginate solution during the experiment (Figure 4.4.5).  Figure 4.4.5: Excess PEI solution cause alginate solution to polymerize. After coating, the microchannel needs to be thoroughly rinsed with water. a) Thoroughly rinsed PEI-coated channel and b) not rinsed PEI-coated channel filled with 1.0 wt% alginate.  Because PEI is positively charged (polar) in nature, contact angle measurement becomes a good indication of the PDMS surface treatment. After the coating procedure, the PDMS surface turns from being hydrophobic to hydrophilic. The immobilization of PEI on PDMS surface may due to the result of van der Waals interaction [117]. The contact angle measurement is shown in Figure 4.4.6. The interaction between PEI and 53  glass surface is expected to be ionic (glass is a negatively charged surface) [118]. Because glass substrate is strongly hydrophilic by default, the same measurement cannot be done with the PEI-coated glass surface.  Figure 4.4.6: The static contact angle measurement shows that the original a) hydrophobic PDMS surface (contact angle 90° – 110°) changes to being b) hydrophilic (contact angle 70° – 80°) after PEI coating.  It is expected that the polymer layer degrades with each gel formation and dissolution process; the surface treatment has to be renewed [44]. Quantitative analyses on the polymer degradation in cell culture media have been tried; there is no observable static contact angle difference between coated and non-coated surfaces. However, a dynamic contact angle difference can be observed even after the PEI-coated PDMS slab is submerged in cell culture media and incubated for 48 hours. Because culture media contains more than 30 nutrients, ions, and proteins, the actual explanation for why no static contact angle difference can be observed remains unknown.  Since all the  microfluidic chips can only be used once, renewing the surface coating is never considered.  54  CHAPTER 5  5 On-Chip Cell Culture 5.1 Biocompatibility Test  5.1.1 Food Dye Coloring For visualizing flow profile, FD&C Blue No.1 and Red No. 40 food coloring were added to the EDTA and CaCl2 solutions. Blue no. 1, also known as Brilliant Blue FCF, is a synthetic food colorant derived from coal tar [119]. The dye is water soluble with maximum absorption at about 630 nm [120]. The dye is certified as a safe additive in the European Union and United States and is often found in dairy products such as ice cream and soft drinks [121]. Red no. 40, which also goes by Allura Red AC, is a water-soluble synthetic food dye. Its maximum absorbance is around 503 nm [122]. Similar to Blue no. 1, Red no. 40 is also derived from coal tar. It is approved by the European Union and by the Food and  55  Drug Administration in the United States for use in puddings, dairy products, beverages, and confections [121]. To ensure that the food coloring did not affect the cell viability during the experiment, a biocompatibility test was carried out using HFL1 fibroblast cells. The experiment set up is shown in Figure 5.1.1 and summarized in the following steps: 1.  Prepared HFL1 cell suspension with concentration of 1 million/ml.  2.  Dispensed 100 ml suspension into each well, resulting in a total of 100,000 cells per well.  3.  Added 1.5 ml of culture medium (Ham’s F12K) in each well.  4.  Added 25 µl of food coloring in each well.  5.  Placed the 6-well plate inside the incubator (5 % CO2 at 37 °C)  6.  Performed cell viability readings every 24 hrs, 48 hrs, and 72 hrs.  Figure 5.1.1: Experimental setup of food dye compatibility test. Each well contained 1.5 ml of culture medium and 100,000 HFL1 fibroblasts. The 6-well plate contained one control group and two experimental groups with 25 µl of Blue No.1 and Red No. 40 food dye.  56  The biocompatibility test ran for three days. The cells were removed from each well and live/dead assays were performed with trypan blue. More information about live/dead assays is discussed in section 5.2.4. The test showed that the food colors used did not affect cell viability (Figure 5.1.2).  Figure 5.1.2: Cell viability reading using trypan blue after cells were exposured to food coloring.  5.1.2 PDMS Curing PDMS is well known as an excellent soft material for soft lithography. It is especially known for its biological usage due to its low curing temperature, fast curing, and non-toxic nature [85, 123]. However, uncured PDMS elastomer is very sticky and viscous to work with.  57  At the beginning of the experiment, the cell viability inside the microfluidic channel was unexpectedly low. We suspected that the PDMS base and curing agent may seep out from not fully cured PDMS sidewalls. To find a proper curing time for PDMS moulding, a series of biocompatibility tests related to PDMS curing were carried out. A sequence of PDMS pallets were made from 6-well culture plates with various curing times at a 50 °C curing temperature. The PDMS was cured at lower temperature than specified by the PDMS datasheet (50 °C instead of 80 °C) to prevent the SU-8 pattern from cracking due to different thermal expansion coefficient between the SU-8 layer and the Si substrate. The curing time of PDMS pallets was from 1-13 hours. The experimental set up was similar to the one carried out with food coloring shown in the previous section. It is described below and shown in Figure 5.1.3: 1.  Prepared HFL1 cell suspension with concentration 1million/ml.  2.  Dispensed 100 ml suspension into each well with a total of 100,000 cells per well.  3.  Added 1.5 ml of culture medium (Ham’s F12K) in each well.  4.  Pre-cured PDMS pallet was plated inside the well.  5.  Placed the 6-well plate inside the incubator (5 % CO2 at 37 °C)  6.  Performed viability every 24 hrs, 48 hrs, and 96 hrs with trypan blue (see section 5.2.4 for more information about viability assay).  58  Figure 5.1.3: Experimental setup of PDMS curing compatibility test. The cured PDMS disks were placed inside a 6-well plate, pre-seeded with 100,000 HFL1 fibroblast cells. Each plate contained 2 wells of control group for comparison.  From the experiment, it was been found that there is a significant drop of cell viability with PDMS at a 50 °C curing temperature when curing time is less than three hours (Figure 5.1.4).  This result gives a good reference for setting up proper  fabrication protocols.  59  Figure 5.1.4: Cell viability reading over four-day period. There is a noticeable difference of cell viability for PDMS when curing is less than three hours at 50°C.  5.2 Experimental Setup For each set of the experiment, six to ten cell-loaded microfluidic chips were prepared. The setup for the experiment had to enable easy access to the device so the cell-loading process could be repeated from chip to chip efficiently. It was expected that the whole cell-trapping process would be completed within one and half hours; otherwise, cell sedimentation and aggregation would affect the distribution of cells inside the channel.  Exposing microfluidic channels to the outside environment for extensive  periods of time would also increase the chance of contamination.  60  5.2.1 Cell Counting The number of cells trapped inside the microfluidic device is highly dependent on the cell concentration of the alginate-medium cell suspension. For cell counting, we used a standard hemocytometer (Bright-Line, Canada).  b)  a)  Figure 5.2.1: Hemocytometer: a) Improved Neubauer hemocytometer. b) Under the microscope view, the hemocytometer has nine large squares. The center square is subdivided into 25 squares and each of these is divided again into 16 squares.  A hemocytometer is a thick glass plate, which fits onto the stage of a microscope. There are several designs of hemocytometers available: Burker, Fuchs-Rosenthal and Neubauer. The one used was the improved Neubauer, the most common type used today (Figure 5.2.1). Under the microscope view, the improved Neubauer-type hemocytometer has nine large squares with 1 mm sides. The center square is subdivided into 25 squares and each of them divided again into 16 0.05 × 0.05 mm squares (Figure 5.2.1). With the cover slip on, the volume of each of the nine large squares is 1 × 1 × 0.1 = 0.1 mm3. After obtaining the average cell count from each of the 9 large squares, we multiplied by 61  1 x 104 to obtain the number of cells per millilitre. The expected error associated with cell counting using the hemocytometer is around 10% [124]. It is desirable to trap around 1000 cells inside the microchannel for viability and drug testing.  For Hep G2 and LCC6/HER2 cell lines, the seeding density is also  adequate for forming 3-D spheroids (section 5.3.2 and 5.3.3). From experience, for trapping 1000 cells inside the gel slab (300 µm wide × 90 µm high × 1 cm long) within the designed microchannel (800 µm wide × 90 µm high  × 1 cm long), 10 million cells/ml of initial alginate cell suspension was needed (Figure 5.2.2).  Figure 5.2.2: Number of trapped cells vs. initial cell suspension concentration. The number of trapped cells in the gel layer (300 µm wide × 90 µm high × 1 cm long) varied linearly with initial cell concentration in the alginate solution. The experiments were carried out with 40 mM CaCl2 solution, 1.0 wt% LF120M alginate solution.  62  Besides the initial cell suspension concentration, the number of cells trapped in the gel layer was also highly dependent on other variables such as cell sedimentation, the formation of cell clumps and alginate gel growth rate. At high gel forming rate, the cell density inside the gel layer would be closer to the initial cell concentration of the alginate cell suspension.  The lower the gel forming rate, the less likely the cells would be  trapped. For the Hep G2 cells, 20 to 25 minutes after the preparation of cell suspension, cells started to sediment and form aggregates. The longer the experiment was carried out, the less evenly distributed was the cell inside the gel layer.  5.2.2 Pressure-driven Fluid Flow Control System A syringe pump or peristaltic pump is often used for applications requiring precision transport of fluid. However, due to the mechanical design of both types of devices, it is difficult to apply gradual changes to independent channels.  Severe  pulsations can also be observed at a low flow rate (nl/min range). To overcome this problem during the cell loading process, the flow of the liquid was controlled by a highprecision pressure regulator (MFCS-8, Fluigent) as shown in Figure 5.2.3. The MFCS-8 has eight independently controlled channels with the smallest pressure step of 25 µBar and a deviation of less than 1%. Pressure can be adjusted from 0 bar to 1 bar.  63  Figure 5.2.3: MFCS-8 high precision pressure regulator  The flow rate inside the microfluidic channel was also strongly dependent on the tubing used. With soft elastic tubing, one may have a slow response time, or at the other extremity, rigid tubing could cause leakage at connection points. From experience, Tygon tubing (0.020” ID, 0.060” OD) was found to be ideal for our purpose (Figure 5.2.4).  64  Figure 5.2.4: Experimental setup during cell-loading process. It was done with the MFCS-8 pressure control device and Nikon TE2000-U fluorescence microscope.  5.2.3 Nutrient Circulation Even though PDMS is highly porous and there is enough oxygen supplied to the cells by simple diffusion [125], there may not be enough glucose or other nutrients to  65  permeating through the PDMS layer as mentioned in section 3.1.3.2. To overcome this problem, a syringe pump was used to continuously supply culture mediums through the micro-bioreactor. To further demonstrate the need for a continuous supply of nutrient through the culture chamber, LCC6/HER2 cells were loaded into four identical microfluidic chips and divided into two test conditions. One set of microfluidic chips was connected to a syringe pump and circulated with DMEM culture medium at a rate of 0.3 µl/min; the other set of microfluidic chips was in the static condition (the channel was pre-filled with medium and there was no circulation provided). All test chips were submerged in the DMEM cell culture medium with inlet and outlet blocked. All microchips were loaded with 200 to 300 cells in alginate gel inside the microchannel. From the experiment, we found that cell viability dropped significantly after 3 days in the static condition, where no nutrient was supplied and no waste was removed (Figure 5.2.5). This result supports the assumption that, without circulation, cells will die due to lack of nutrients and accumulation of waste generated from metabolism.  66  Figure 5.2.5: Cell viability comparison between the static condition and perfusion condition (continuous supply of nutrient at a rate of 0.3 µl/min). We found that, without circulation, cell viability dropped significantly as predicted.  Besides the above reasons, perfusion cell culture also provides a more stable culture environment due to the continuous supply of nutrients and removal of waste. Continuous perfusion, in combination with 3-D culture with multiple cell types, can provide a microenvironment that mimics the structure and composition of a blood vessel [40, 126]. This brings a whole new dimension to drug testing. For circulating the fluid, the Tygon tubing purchased from Fisher Scientific was connected to the pre-drilled access holes on the PDMS chip. The syringe pump was set at the withdraw motion to prevent contamination from the tubing and provide easy access to waste removal. Since fresh mediums never came in contact with the tubing for extended periods of time, the biocompatibility of Tygon tubing was not a concern. The  67  size of the syringe was chosen to be as small as possible to minimize the pulse flow generated by the syringe pump (Figure 5.2.6).  Figure 5.2.6: Syringe pump was used to continuously supplied culture media to the chip. The pump was set at the withdraw motion. The chips were placed inside the incubator to maintain a stable microenvironment.  Glucose is the major energy source for cell metabolism and the main component of all culture medium. The flow rate of the culture medium was determined by the rate of cell glucose consumption. Based on literature research, the average glucose consumption rate for mammalian cells is ~5 × 10-4 µg/103cells/min [127]. In DMEM, the glucose concentration is 4500 mg/L. When the DMEM culture medium was flowing through the channel at the rate of 0.3 µl/min, as it was in the experiment, the glucose was supplied at  68  a rate of 1.35 µg/min. If we assume 1000 cells were seeded inside the channel, the glucose supply rate was more than three orders higher than the rate of average cell consumption. The syringe pump was set at a higher withdraw rate than the average nutrient consumption rate needed by the cells to ensure the continuous supply of culture medium (Channel blockage and tubing leakage may occur during long-term cell culture experiments).  5.2.4 Live/Dead Assay Viability is a measure of the metabolic state of a cell population, which indicates the proportion of live, metabolically active cells in a culture. The cell viability may be determined from simple assays such as dye exclusion where the loss of viability is determined by membrane damage [124].  5.2.4.1 Trypan Blue For cell viability testing, trypan blue is one of the most common vital stains used. The assay is derived from toluidine, that is, any of several isomeric bases, C14H16N2. It has a very deep, blue color. Live cells or tissues with intact cell membranes will not be colored; however, the dye will traverse the dead cell membrane. As a result, the dead cells appear as a distinctive dark-blue color under white light. Trypan blue dye only stains the dead cells and excludes the live cells. This is also called the Dye Exclusion Method. It is fast and effective.  69  The dye is very stable under room conditions. For taking the reading, an equal volume of trypan blue was added to a cell suspension. The dye is poisonous. Therefore, the result had to be interpreted within 2 to 3 minutes; after that, all the cells would begin to take up the dye [124].  5.2.4.2 Live/Dead Viability/Cytotoxicity Kit Trypan blue gives instant result, but the samples can only be interpreted once, which is extremely inefficient. In a later stage of the experiment, a new assay from Invitrogen was used. The live/dead kit from Invitrogen contained two color dyes, calcein AM and ethidium homodimer-1 (EthD-1). Calcein AM (ex ~495nm, em ~515nm) was retained within live cells and EthD-1 (ex ~495nm, em ~635nm) was excluded by the intact plasma membrane of live cells [128]. Live cells were identified by the presence of ubiquitous intracellular esterase activity, which turns the non-fluorescent cell-permeate calcein AM to an intensely fluorescent calcein. In this case, the live cell showed a green fluorescence color and the dead cell nucleolus showed a red fluorescence color [128]. Unlike trypan blue, this assay is not lethal to the cell. The sample can be re-dyed in the future, as necessary. For dying the microfluidic chip, 4 µM of EthD-1 was prepared with PBS and 2.5 µM of calcein AM was mixed right before the experiment. The reagent was injected into the channel with a syringe and incubated for 45 minutes. The reagents needed to be stored below -20°C. Calcein AM is susceptible to hydrolysis when exposed to moisture, so the solution needed to be freshly prepared  70  before use. EthD-1 is stable in a moisture environment. A stock EthD-1 solution was prepared with PBS beforehand, which has a shelf life of up to one year under frozen conditions [128].  5.3 Cell Viability  5.3.1 HFL1 Fibroblast Cells Fibroblast is a type of cell that synthesizes the extracellular matrix and collagen, the structural frameworks for animal tissues. connective tissues in animals.  It is the most common cell within  The main function of fibroblasts is to maintain the  structural integrity of connective tissues [33]. Fibroblasts are among the most widely used cells in laboratory cultures due to the relative ease in maintaining them and low bio-safety level [124]. To first test out the reliability of this culture technique, we trapped ~300 human HFL1 fetal lung fibroblasts (ATCC, USA) in the microchannel. For comparison, the same cell line was also cultured inside alginate gel beads (350 µm diameter) and in a typical culture flask as control groups (Figure 5.3.1).  71  Figure 5.3.1: HFL1 cell viability comparison. The cell viability inside the microchannel was as high as cell culture inside the Petri dish and alginate gel beads.  For culture HFL1, we used Ham's F12K medium (Invitrogen USA) supplemented with 10% Fetal Bovine Serum (FBS), 100 units of Penicillin and 100 µg of Streptomycin. For cell culture inside the microchannel, Ham’s F12k medium was continuously fed through the device. The setup of the experiment is shown in section 5.2.3. The syringe pump was set with a withdrawing rate of 0.3 µl/min. The cell viability is observed every 24 hours for 2 days. From the results, we proved that the cell viability inside the channel was as good as other cell culture techniques (Figure 5.3.2).  72  b)  a)  c)  Figure 5.3.2: HFL1 cellular shape: a) In a typical culture flask, HFL1 extended out and adhered to the bottom of the plate. b) The HFL1 cells were in spherical shapes inside the alginate beads. c) Similar spherical structures also appeared inside the microchannel,  5.3.2 Hep G2 Hypatocyte Cells The Hep G2 cell line was originally derived from human liver tissue [129]. The cells secrete a variety of major plasma proteins (e.g. albumin, alpha2-macroglobulin, alpha 1-antitrypsin, transferrin and plasminogen), which are often used as an indicators for studying metabolic functions [33, 130] since liver is the main organ associated with drug detoxification. To explore drug metabolism and toxicological testing, a cell culture that simulates in vitro models of liver is required [129].  73  Isolating hepatocytes using 2-D culture techniques has posed a great challenge based on the fact that the cell line often shows loss of viability and metabolism activity [131, 132]. When culture hepatocytes using hydrogels, the cells form spheroids and possibility retain expressions of liver markers (such as albumin) and their ability to respond to peroxisome proliferators [132]. It is a more stable, attractive tool for further investigation of the acute phase and other hepatocellular responses. To further improve the in vitro model, we demonstrated the possibility of 3-D culture the Hep G2 cell line inside the microdevice. The hepatocyte cell line was cultured inside the channel for 2 weeks. Hepatocytes are much smaller than fibroblasts in terms of cell size. They also grow much faster. For the experiments, we managed to trap 500 to 1000 cells inside a 300 µm wide x 90 µm high x 1 cm long gel slab. For the Hep G2 culture, the growth medium used was Minimal Essential Media α (MEM α) (Invitrogen, USA) supplemented with 10% FBS, 100 units of Penicillin and 100 µg of Streptomycin. The setup was the same as the one used for the HFL1 fibroblast in section 5.3.1. The culture medium flowed flowing through the microchip at a rate of 0.3 µl/min. For comparison, we cultured the same batch of cells in 350 µm alginate beads and in a culture flask along with the cells in the alginate gel layer inside the channel (Figure 5.3.3).  74  b)  a)  c)  Figure 5.3.3: Hep G2 cellular shape: a) In a typical culture flask, the Hep G2 cell line only formed a 2-D structure at the bottom of the plate. b) After 6 to 7 days, the HepG2 cell line formed spheroids inside the 350 µm alginate beads. c) Similar 3-D spheroids were also observed inside the microchannel after 6 days.  We observed that the Hep G2 started to form spheroids in the gel beads and microchannel on the 5th day of the experiment. As time went by, the cell proliferated and the spheroids grew in size (Figure 5.3.4).  75  a)  b)  c)  d)  Figure 5.3.4: The Hep G2 cell line inside the microchannel forms 3-D spheroids. Pictures were taken throughout the experiment. a) The first day of the experiment. b) 1 week into the experiment. c) 2 weeks into the experiment. d) At the end of the experiment, the cells were dyed with the live/dead kit. The result shows that more than 95% of the cells were alive.  The experimental results are shown in Figure 5.3.5. From the results, we can say that, for long-term culture, the cell viability inside the channel is as high as traditional 2D culture flask and 3-D alginate beads culture techniques.  76  Figure 5.3.5: Hep G2 cell viability comparison. The cell viability inside the microchannel is as high as cell culture inside a petri dish and alginate gel beads.  5.3.3 LCC6\HER2 Breast Cancer Cells Malignant neoplasm, known by most people as cancer, is a disease in which a group of cells display uncontrolled growth (division beyond the normal limits), invasion (intrusion onto the adjacent tissues), and sometimes metastasis (spreading to other locations in the body via lymph or blood). According to the American Cancer Society, more than 7.6 million people in the world died from cancer during 2007 and breast cancer is the number one cause of death for female cancer patients [133, 134]. There is great interest in cancer drug development. To make the microfluidic device feasible for drug testing, we first needed to demonstrate that it is feasible to culture cancer cells inside the microdevice for extensive periods of time. In this case, we  77  used LCC6/HER2 breast cancer cells. Similar to hepatocytes, breast cancer cells also formed spheroids inside the gel matrix after 5-7 days (Figure 5.3.6).  b)  a)  c)  Figure 5.3.6: LCC6/HER2 cellular shape. Similar to hepatocyted a) the LCC6/HER2 cell line only formed a 2-D structure at the bottom of the plate. b) After 5 to 7 days, the LCC6/HER2 cell line formed spheroids inside the 350 µm alginate beads. c) 3-D spheroids also formed inside the microchannel.  We cultured the LCC6/HER2 cancer cell line for 3 weeks. Starting on the 6th day, the cancer cells started to form spheroids.  Throughout the experiment, the cells  proliferated and the spheroids grew in size (Figure 5.3.7).  78  a)  b)  c)  d)  e)  Figure 5.3.7: The LCC6/HER2 cells proliferated and formed 3-D spheroids inside the gel layer. Pictures were taken throughout the 3-week experiment. a) The first day of the experiment. b) 1 week into the experiment. c) 2 weeks into the experiment. d) 3 weeks after the start of the experiment. e) At the end of the experiment, the cells were stained with the live/dead kit. The result shows that more than 95% of the cells were alive.  79  The experimental results are shown in Figure 5.3.8. The cell viability inside the microchannel was consistently over 90% throughout the 21 days.  In terms of cell  viability, the 3-D microchannel culture technique was as reliable as the typical culture technique in the culture flask or 3-D culture inside alginate gel beads. Based on this result, we could precede to cancer drug screening (section 5.6).  Figure 5.3.8: LCC6/HER2 cancer cell viability comparison. The tumor cells were cultured inside a microdevice for 3 weeks. The result shows that the cell viability was more than 90% consistently throughout the experiment.  5.4 Co-culture By manipulating the fluid flow, we achieved separate gel formation at two boundaries close to the channel sidewalls or multiple stacked adjacent layers. This allowed different cell types to be cultivated in close proximity in the same cell culture environment (see Figure 5.3.9, Figure 5.3.10).  80  a)  b)  c)  Figure 5.3.9: The trapping process for co-culture two cell types simultaneously. a) Different cell types were mixed in two separate alginic acid solutions. We used the interior and central inlets for EDTA and CaCl2. This was the initialization step. A laminar flow of EDTA was used to separate the Ca2+ and alginate flows prior to gel formation. b) After the EDTA flow was suppressed, the Ca2+ ion solution came into contact with the alginate phase. The gel started to grow from the boundary towards the alginate stream with cells trapped inside. c) Once the gel formation was complete, the inlet flows were switched to culture medium for long-term cell culture.  81  b)  a)  Figure 5.3.10: Co-culture HFL1 and Hep G2 cells. Cells were individually cultured inside microfluidic devices: a) two separate regions or b) adjacent layers. Cell viability was measured using a LIVE/DEAD Viability/Cytotoxicity Assay Kit immediately after gel formation. The live cells showed an intense uniform green fluorescence.  5.5 Cell Releasing To demonstrate the possibility of releasing cells from the microchip, we first cultured 1000 to 2000 Hep G2 cells inside the microfluidic device for 48 hours and gently injected 40mM EDTA solution to dissolve the alginate gel. Two control groups were prepared side by side with this experiment. One had Hep G2 cells cultured inside typical culture flasks. Another set of Hep G2 cells were trapped inside alginate beads with diameters from 350 µm to 1 mm. A 0.5 cm diameter reservoir was created at the end of the channel to accommodate the dissolved cell suspension. The cell suspension was then pipetted out into a 12-well plate and cultured for 5 days (Figure 5.3.11).  82  a)  b)  Figure 5.3.11: Cell releasing: a) To retrieve the cells trapped inside the microfluidic device, a 0.5 cm reservoir was created at the end of the channel to accommodate the released cell suspension. b) The majority of Hep G2 Cells remained alive 5 days after releasing.  We found that cell viability was much higher with the one released from the channel than the one released from alginate beads (Figure 5.3.12). From cell releasing experiments using alginate beads, it has been found that the cell viability is highly dependent on the concentration of EDTA solution used and the length of time cells are in contact with the EDTA solution. The longer the cells are in contact with the EDTA solution and the higher the EDTA solution concentration used, the lower the cell viability. It only takes 15 to 20 seconds to release the cells from the channel using 40 mM EDTA solution, but it will take 1 hour to 1.5 hours to release the cells form 1 mm 1.0 wt% alginate gel beads. This shows another advantage of culture cells inside the microchannel.  83  Figure 5.3.12: Cell releasing viability comparison. The cell viability was noticeably higher when releasing from the microchannel than releasing from the beads due to the shorter exposure time to EDTA.  5.6 Drug Testing Unfortunately, oncology has one of the poorest records for investigational drugs in clinical development, with success rates that are more than three times lower than those of cardiovascular disease [135]. Anticancer drug targets can be classified into two categories: essential and non-essential. Essential means that at least one vital cell type in the body depends on the target for survival. Drugs that target essential functions most likely have very narrow therapeutic windows. Drugs that target essential genes always have the risk of serious side effects. On the contrary, our body has much higher tolerance for drugs that target non-essential proteins. However, the efficacy of non-essential drugs is limited. Another challenge with drug treatment is that essential and non-essential genes are very difficult to discriminate within mammals [135].  84  Cancer drug development faces many challenges.  However, microfluidic  technologies that can carry out tests based on a single cell or have access to a more detailed picture of cell abnormalities bring new possibilities to drug development. For the experiment, we used doxorubicin (Dox) generously provided by Dr. Helen Burt, UBC Faculty of Pharmaceutical Sciences.  5.6.1 Doxorubicin (Dox) Doxorubicin (Dox), also known as adriamycin or hydroxydaunorubicin, has a chemical formula of C27H29NO11 (Figure 5.3.13) and MW of 543.52 g/mol. Dox has peak excitation and emission wavelengths at 480 nm and 550 nm respectively. Dox is a common chemotherapy drug used in treatment for many types of cancer [136, 137]. For chemotherapy, Dox is normally given by injection into a vein through a canula. The treatment is given in sessions over a few months. Possible side effects may include hair loss, nausea, vomiting, and lowered resistance to infection. The exact mechanism of doxorubicin is complex and still somewhat unclear. What is known is that it interacts with DNA by intercalation and inhibits biosynthesis [137]. It also has been reported to interact directly with the cell membrane and to inhibit the unwinding of DNA for replication [138]. This phenomenon would inhibit cell growth.  85  Figure 5.3.13: The molecular structure of doxorubicin (Dox).  The ability of doxorubicin to interfere with the growth of LCC6/HER2 breast cancer cells was later determined through both fluorescence and confocal microscopes (section 5.6.4).  5.6.2 Drug Diffusion in Alginate Gel Lacking any non-invasive techniques to measure the concentration of Dox inside the microfluidic device during drug testing, a low concentration of the fluorescent dye, Rhodamine B, was introduced to evaluate cross channel diffusion. Rhodamine B is a commonly used hydrophobic fluorescent dye in biotechnology applications such as fluorescence microscopy, flow cytometry, and ELISA. The chemical structure of the fluorescent dye is shown in Figure 5.3.14. The peak excitation and emission wavelength of Rhodamine B dye are 540nm and 625nm [139].  With the chemical formula,  C28H31ClN2O3, and molecular weight of 479.01 g/mole, it is comparable with Dox in terms of molecular size. We used Rhodamine dye to simulate Dox for measuring the diffusivity because it is less toxic and safer to use.  86  Figure 5.3.14: Structures of Rhodamine B (C28H31ClN2O3) [140].  The intensity of the Rhodamine B dye in the channel was used to correlate with the dye concentration, which represented the drug concentration (Figure 5.3.15).  87  Figure 5.3.15: Diffusion time measurement of Rhodamine B through alginate gel layer. A 200-µm gel layer was formed inside the channel. 500 µM Rhodamine B was injected at a rate of 3 µL/min into the channel at room temperature to simulate Dox (cancer drug). The intensity of the fluorescence signal can be used to determine the dye concentration to show the amount of dye diffusing into the gel layer across the channel. Since Rhodamine B has a similar diffusion coefficient as Dox, it is a close approximation to the delivery of cancer drugs to the breast cancer cells.  88  To estimate the diffusion coefficient of Rhodamine B, we can use Fick’s second Law:  ⎧ x ⎫ c( x, t ) = c0erfc ⎨ ⎬ ⎩ 2 Dt ⎭  (5.1)  c( x, t = 0) = 0  (5.2)  c( x = 0, t ) = c0  (5.3)  c ( x = ∞, t ) = 0  (5.4)  With boundary condition:  where erfc(q) is known as the complementary error function, which can be written as:  erfc(q) =  ∞  2  π  −t ∫ e dt 2  (5.5)  q  With the channel height equal to 90 µm, Gel width equaled to 200 µm, and original Rhodamine B concentration equal to 500 µM. Based on the measured data (Figure 5.3.15), the diffusion coefficient in alginate gel ( ≈ 5.0 × 10−10 m 2 / s ) at room temperature, which is very similar to the diffusion coefficient of Rhodamine B in aqueous solution ( ≈ 3.6 × 10 −10 m 2 / s ) at 21.5°C [140].  5.6.3 Drug Diffusion through PDMS Chip To have precise control over the drug testing condition, it is necessary to ensure that tumor cells only in contact with the Dox solution passing through the channel. Because the PDMS chip is porous and submerged in the Dox solution, we need to make certain that Dox does not diffuse through the chip and reach cells inside. Similar to  89  section 5.6.2, we replace Dox with Rhodamine B for measuring the diffusion rate of Dox though PDMS. For determining the diffusion speed, a PDMS pallet is submerged in 500 µM Rhodamine B solution. The diffusion distance (the distance Rhodamine B diffuse from the edge of PDMS block towards the center of the PDMS pallet) is measured through fluorescence microscope for 2 weeks (Figure 5.3.16).  Because of the hydrophobic  nature of Rhodamine B, hydrophobic attraction is expected between the dye and the PDMS bulk [141].  Figure 5.3.16: Diffusion distance of Rhodamine B through PDMS block over two weeks at room temperature.  Using Fick’s second law of diffusion (5.1), we estimate the diffusion coefficient D ≈ 2.5 × 10−11 m 2 / s . Base on the geometry of the designed microfluidic device, it will take more than four days for the fluorescent dye to reach the main section of the channel.  90  5.6.4 Control Group To ensure that all the nutrient consumed by the cells inside the microfluidic device was provided by the withdrawing motion of the syringe pump, four different control groups were test: submerged in culture medium flowing with culture medium, submerged in culture medium flowing with PBS, submerged in PBS flowing with culture medium, and submerged in PBS flowing with PBS. The microfluidic chip was placed inside the six-well plate with fluid drawn from a separate well through PTFE tubing (Figure 5.3.17). Tygon tubing is more flexible and easier to use, but biocompatibility is always a concern. For the case where supplements were required to flow through, PTFE tubing was used. For the four different control cases, cell viability was observed after 72 hours using the Live/Dead kit purchased from Invitrogen. The result showed that with the continuous feeding of nutrients, the cell viability remained stable and above 90%. With the two cases of feeding only with PBS, the cell viability dropped significantly below 5% (Figure 5.3.18).  91  Figure 5.3.17: Control experiment before drug testing. The microfluidic chip was submerged in culture medium or PBS with culture medium or PBS flowing through from a separate well connected with PTFE tubing. In this case, the cell culture chip was submerged in culture medium with PBS flowing through.  92  Figure 5.3.18: Control experiment result. The cell viability remained above 90% for the two cases with the continuous feeding of the nutrients. For the two cases involving feeding with only PBS, the cell viability dropped below 5%. The result was taken after 72 hours.  5.6.5 Experimental Result For drug testing, the Dox solution was prepared with 0.2% DMSO and DMEM culture medium. The cells were first loaded in the channel and incubated for 24 hours with a circulation of drug-free culture media. After 24 hours, the drug-free culture media was replaced with 0.8 µM and 1.5 µM of the Dox solution. The drug solution was continuously perfused through the channel at a rate of 0.3 µl/min. The Dox concentration (0.8 µM and 1.5 µM) was previously determined from the MTS assay, where the cells were encapsulated in 50 µl 1.0 wt% alginate gel beads. Each gel bead contains about 2500 cells. The IC 50 for Dox with the LCC6/HER2 cell line in  93  alginate beads was determined to be 0.4 µM. At this concentration, the drug was 50 % effective in inhibiting cell proliferation. From drug encapsulation research, it has been found that alginate (negatively charged) have a strong ability to accumulate positively charged Dox [142, 143]. For this reason, the drug concentration used was 2 to 3 times higher than the IC 50 (0.8 µM and 1.5 µM) to insure enough Dox can reach the tumor cell. Two control groups were prepared: one with the typical DEMEM culture medium and the other one feeding with DEMEM and 0.2% DMSO (all drug solutions were prepared with 0.2 % DMSO). Each channel contained 500 to 1,000 cells. The set up was the same as the one mentioned in section 5.2.3. Cell viability readings were taken every 24 hours for 3 days after introducing the cancer drug (Figure 5.3.19).  Figure 5.3.19: Cell viability reading for Dox drug testing. The viability reading was taken every 24 hours for 3 days after introducing 0.8 µM and 1.5µM Dox to the cells.  94  From the results, we found that cell viability gradually lowered after 48 hours. Dox did not kill the cell on contact but stopped the cells from proliferating. Therefore, the effect of the drug can only be observed after 24 hours. Additional pictures were taken that show that the LCC6/HER2 cell line formed spheroids in the control chip with 0.2 % DMSO, but the same structure could not be observed after contact with 0.8 µM Dox (Figure 5.3.20, Table 5.3-1). The experimental procedure is shown in Appendix D.  a)  Without Dox (control)  b)  3 days with Dox (test)  Figure 5.3.20: Drug testing result: a) The LCC6/HER2 cell line formed spheroids in the control chip with 0.2 % DMSO 5 days into the experiment. b) Dox stopped the cells from proliferation so 3-D spheroid structures could not be observed after contact with 0.8 µM Dox.  95  Table 5.3-1 Cell proliferation after 72 hours treatment.  Increase in Cell Number  Control  0.8 µM Dox  1.5 µM Dox  >150%  ~0%  ~0%  96  6 CHAPTER 6  Conclusion 6.1 Summary In this project, we have designed a microfluidic device, which is capable of reversible trapping of mammalian cell lines using alginate, ionic cross-linking hydrogel, in combination with Ca2+ ions. In the microfluidic device, cells are cultured in ECM-like structures. With alginate gel trapping, nutrients had access over the whole surface of the cell and the immobilization process occurred under mild conditions at room temperature. Because the microfluidic device is made with PDMS and glass (transparent to most of the fluorescent marker), it is compatible with many biological imagining techniques. In order to provide enough nutrients to the cultured cells and remove metabolic waste, culture media was continuously fed to the microchip with a syringe pump to provide a stable microenvironment for long-term cell culture.  We have also  demonstrated the possibility of co-culture two cell types (HFL1 and Hep G2) in separate gel layers. Continuous perfusion, in combination with 3-D culture with multiple cell 97  types, can provide a microenvironment that mimics the structure and composition of a blood vessel. Various mammalian cell lines were successfully cultured within a microfluidic device for long periods of time. The HFL1 cells have successfully cultured for over 48 hours. The Hep G2 and LCC6/HER2 cells were cultured for more than 2 and 3 weeks with cell viability of more than 90%. After culture the cells for more than 5 days, hepatocytes (Hep G2) and breast cancer cells (LCC6/HER2) formed 3-D spheroids. The 3-D structures have a greater resemblance to the cell behaviour in the in vivo conditions and could only be observed when the cell lines were cultured within the gel matrix. In the last part of the project, we demonstrated the possibility of performing cancer drug testing within the microfluidic device. LCC6/HER2 breast cancer cells were first cultured within the microdevice for 48 hours. 1.5 µM and 800 nM of the cancer drug, Dox, were introduced to the cells. Dox inhibited the unwinding of DNA thus stopping the cells from proliferation. Dox did not kill the cell on contact so the effect of the drug could only be observed after 24 hours. The observed result agrees with the prediction.  6.2 Future Work Current experimental results are limited to cell viability reading and manual cell counting. Due to the dimension of the designed device it is nearly impossible to perform conventional MTS assays using a plate reader. Future work would focus on applying MTS or similar assays within the microfluidic device to investigate cellular activities during Dox drug testing.  98  MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) is a colorimetric assay for viable cells. The dye is cleaved to a colored product by the activity of dehydrogenase enzymes and this indicates levels of mitochondrial activity in cells. The color development is proportional to the number of metabolically active cells [124, 144]. It is one of the most commonly used assays to evaluate cellular activity. 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Cells were subject to shear stress.  None  • One of the earliest cellular study using microchannel. Insect cells were cultured with in static microfluidic environment [38].  None  • Long term culture inside microenvironment for further study [31]. Cells were subject to shear stress.  None  • Cells were cultured in static microenvironment to characterize the device which generates a precise dose of dissolved oxygen [148].  None  111  Bacillus subtilis bacteria  Embryos / Oocytes  Rate hippocampal nerve  B35 rat neuroblastoma  Fibroblast (CRL10225, ATCC)  CHO Epithelial  Hepatocytes (Hep G2)  • Bacillus subtilis were cultured within static microenvironment to demonstrate dynamic aerotaxis assays driven by oxygen microgradients [148].  None  • One of the earliest cellular study using microchannel. Manipulate and culture embryos and oocytes to improve efficiencies in assisting reproduction [149]. Cells were subject to shear stress when changing media.  None  • Hippocampus was cultured to form neural networks for biological studies [150]. Cells were subject to shear stress when changing media.  None  • Neuroblastoma cells were cultured to prove the microfluidic chip coated with parylene C could sustain cell growth [151]. Cells were subject to shear stress when changing media.  None  • Mechanical sieves were used [152]. Cells were subject to shear stress during trapping and culturing.  None  • Monitor the cell activities inside Nanoliter Bioreactors [152]. Cells were subject to shear stress. • CHO cells were immobilized for in situ monitoring of the calcium flux response to agonists [153]. Cells were subject to shear stress.  None  • Studies oxygen and nutrient consumption , and the synthesis of albumin and urea in microchip environment [26, 27]. Cells were subject to shear stress. • Test for drug screening and toxicology [154]. Cells were subject to shear stress.  None  112  • Cells were trapped with mechanical sieves [152]. • Multiple drug gradient generator is integrated for drug discovery application [155]. Cells were subject to shear stress. • Demonstrate the possibility of cell culture in microchannel [29]. Cells were subject to shear stress.  None Rat Hepatocytes (AML-12)  • Cells were cultured in a microfluidic device that was made of agarose gel [52]. Cells were in contact with 70°C agarose solution.  Agarose  E Coli  Human promyelocytic leukemia cells (HL60)  LADMAC / BAC 1.2F5  • Study E Coli cell growth rate and cell division time in a single cell culture environment [156]. • A steady gradient of α-methyl-DLaspartate (attractant) was applied to carry out chemotaxis experiment [157]. Cells were subject to shear stress.  None  • A steady gradient of formyl-MetLeu-Phe (f-MLP, attractant) is applied to carry out the chemotaxis experiment [157]. Cells were subject to shear stress. • HL-60 cell line is used for studying chemotaxis that is induced by chemoattractants [158].  None  • Separate microchannel using matrigel to achieve co-culture (LADMAC and BAC cells) [159]. Cells were subject to shear stress.  None  • Co-culture these two cell types to produce an in vitro model of zonation [154]. Cells were subject to shear stress.  Hepatocyte / None Fibroblast  113  poly(ethylene glycol) (PEG)  Human primary mesenchymal stem cells (hMSCs) Fibroblast (IMR-90) / Vein smooth muscle cells (HUVSMC) / Vein endothelial cells (HUVEC)  Pluronic F-127  Collagen, Collagenchitosan, matrigel and fibrin  Yeast cell  Alginate  HEK293 (CRL-1573, ATCC)  Alginate  U-2 OS  • Demonstrate the possibility of culture hMSCs in an automated microfluidic device and monitor the effect of osteogenic differentiation factors on cell motility [160]. • Co-culture multiple cell types together to form layer-by-layer tissue like 3-tunic structure [161]. Cells were subject to low temperature (0°C) • Yeast cells were cultured to demonstrate the possibility and versatility of reversible cell trapping using alginate [44, 162]. • Encapsulated in microbeads inside microdevice [163]. • Demonstrate the possibility of encapsulating cells inside microbeads [163].  Alginate  • Study the function of cells inside alginate microbeads [163].  Alginate Adrenal pheochromocytoma (PC12)  • Perform cytotoxicity assays to the cultured cell line and demonstrate the possibility of co-culture more than one cell type [25]. Cells were exposed with UV light  • Nerve growth factor is introduced in the microchannel through microvalves to study the cell differentiation and axon elongation [164].  None  • Compare the metabolic activity  Chondrocyte  (lactate production rate) and matrix (glycosaminoglycan) production of chondrocytes between microscale and larger culture scale [86]. Cells were in contact with agarose  Agarose  114  solution at higher temperature.  • The beating state of cardiomyocytes is studied for drug (Haloperidol) testing [165]. Cells were cultured in static condition.  Cardiomyocytes  Agarose  Human lung epithelial (A549, ATCC)  Aldehydes + intercellular polymeric linker  Human liver cell line (C3A, ATCC)  Aldehydes + intercellular polymeric linker  Rat primary progenitor cells (BMSCs)  Aldehydes + intercellular polymeric linker  115  • Cells were modified with aldehydes to demonstrate the possibility of gelfree 3-D culture [166]. • After modified with aldehydes, cell clumps were formed without the use of gel encapsulation [166]. • Demonstrate the possibility of 3-D culture without the use of hydrogel after modify the cell surface with aldehydes [166].  B. Chip Fabrication Recipe  116  C. Mammalian Cell Suspension Preparation Protocol 1.  Suck out / Pipette out the media (make sure the needle does not touch the plane of the flask with cells, suck out / pipette out the liquid by tilting the flask - thus all the liquid is in the corner)  2.  Wash cells with PBS (3ml)  3.  Suck out / Pipette out PBS  4.  Add Trypsin + EDTA (2ml)  5.  Bang flask on all 4 sides to loosen up the cells  6.  Incubate the cells for 2-3 minutes and check under microscope to see if all the cells have detached (i.e. all the cells will appear to be moving and floating on the liquid)  7.  Add 2 ml of culture media (+FBS) to the flasks  8.  Pipette up and down for the at lest 10 times – this ensures cells are evenly suspended and there are no clumps  9.  Transfer the cells into a 15 ml conical tube  10.  Centrifuge the tube for about 5 min  11.  Discard the supernatant  12.  Re-suspend the cells in about 2 ml of media – pipette up at least 10 times to ensure even suspension  13.  Take out 10µl of the cell suspension and count under microscope using a Hemocytometer, calculate the concentration of cells and amount of cell suspension needed for the experiment  14.  Make the required suspension – diluting or concentrating the stock (if necessary)  117  D. Drug Testing with LCC6/Her2 and Doxorubicin Control: Cells in chip + gel + media (no DMSO, no drugs) Cells in chip + gel + media + DMSO (no drugs) Cells in wells + no drug Cells in wells + DMSO (no drug) Cells in wells + drug Cells in wells + gel + no drug Cells in wells + gel + DMSO (no drug) Cells in wells + gel + drug Test: Cells in chip + gel + drug Day 0: All the cells loaded Control chip dyed Collect the cells from culture flask Cell count Make up the cell concentration Initial cell viability test (with Trypan blue) Load the cell in the test and control chip + beads + wells Day 1: Overall check up of the wells and the chips + dying one of the control chips (Live/Dead kit) for initial viability test Day 2: Put in the drug (0.8 µM and 1.5 µM) Day 3: One (or two) of the chips with the drug to be dyed (Live/Dead kit) for 24 hr viability test + One of the control chips Day 4: One (or two) of the chips with the drug to be dyed (Live/Dead kit) for 48 hr viability test + One of the control chips (one not dyed before) Day 5: One (or two) of the chips with the drug to be dyed (Live/Dead kit) for 72 hr viability test + One of the control chips (one not dyed before)  118  

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