Microfluidic Studies of Biological and Chemical Processes

Ethan Tumarkin

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Chemistry University of Toronto

Ethan Tumarkin

Doctor of Philosophy

Department of Chemistry University of Toronto

2012 Abstract

This thesis describes the development of microfluidic (MF) platforms for the study of biological and chemical processes. In particular the thesis is divided into two distinct parts: (i) development of a MF methodology to generate tunable cell-laden microenvironments for detailed studies of cell behavior, and (ii) the design and fabrication of MF reactors for studies of chemical reactions.

First, this thesis presented the generation of biopolymer microenvironments for cell studies. In the first project we demonstrated a high-throughput MF system for generating cell-laden agarose microgels with a controllable ratio of two different types of cells. The MF co-encapsulation system was shown to be a robust method for identifying autocrine and/or paracrine dependence of specific cell subpopulations.

In the second project we studied the effect of the mechanical properties on the behavior of acute myeloid leukemia (AML2) cancer cells. Cell-laden macroscopic agarose gels were prepared at varying agarose concentrations. A modest range of the elastic modulus of the agarose gels were achieved, ranging from 0.62 kPa to 20.21 kPa at room temperature. We observed a pronounced decrease in cell proliferation in stiffer gels when compared to the gels with lower elastic moduli.

The second part of the thesis focuses on the development of MF platforms for studying chemical reactions. In the third project presented in this thesis, we exploited the temperature dependent

solubility of CO2 in order to: (i) study the temperature mediated CO2 transfer between the gas and the various liquid phases on short time scales, and (ii) to generate bubbles with a dense layer of colloid particles (armoured bubbles).

The fourth project involved the fabrication of a multi-modal MF device with integrated analytical probes. The MF device comprised a pH, temperature, and ATR-FTIR probes for in- situ analysis of chemical reactions in real-time. Furthermore, the MF reactor featured a temperature controlled feedback system capable of maintaining on-chip temperatures at flow rates up to 50 mL/hr.

Key words: microfluidics, microenvironments, high-throughput, encapsulation, co-culture, biopolymers, elastic modulus, multi-modal, modular, integrated analytical probes, titrations

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Imagination is more important than knowledge. Knowledge is limited.

- Albert Einstein

Acknowledgments

First and foremost I want to give my outmost gratitude to my supervisor, mentor, and friend, Professor Eugenia Kumacheva. Without a doubt, her guidance, wisdom, enthusiasm, and constant support have shaped me into the person, teacher, and researcher that I am today. I am forever grateful for her insights about science and life.

I am also extremely grateful to my committee members Professor Axel Guenther, Professor Gilbert Walker, and Professor Aaron Wheeler.

In addition, I want to give my utmost appreciation to my collaborators Professor Axel Guenther, Professor Gilbert Walker, Professor Peter Zandstra, Professor Barbara Sherwood Lollar, Professor Mark Ungrin, Professor Zhihong Nie, and Professor Jesse Greener for their support and helpful discussions over the past 4 years.

I want to thank all the persons I have had the pleasure of interacting with since I was a young undergraduate summer student during the summer of 2005. In no particular order, I want to say thank you to Dr. Hong Zhang, Dr. Mallika Das, Ilya Gourevich, Dr. Zhixiang Wei, Dr. Alla Petukhova, Dr. Chantal Paquet, Dr. Wei Li, Dr. Minseok Seo, Dr. Lindsey Fiddles, Andrew Paton, Dr. Ryan Simms, Michael Debono, Dr. Daniele Fava, Neta Raz, Dr. Raheem Peerani, Dan Voicu, Lsan Tzadu, Milad Abolhasani, Pasquale Benvenuto, Jörg Fochtmann, and Dr. Jemma Vickery.

In the past 7 years I have had the pleasure of working with people which have become like family to me. Dr. Anna Lee, Yannick Bohren, Dr. Jai Il Park, Dr. Diego Velasco, Dr. Kun Liu, Dr. Slava Dubinsky, Miguel Neves, and Ivan Gorelikov have been like family watching over my shoulders and stepping in during difficult times. My life has been enriched because of them.

Finally, I want to give thanks to my family. To my sister, I want to say thank you for always being supportive and helpful. Always know that I am very proud of you. To my father, I want to say thank you for your unparalleled belief in me, for your constant curiosity and willingness to listen to my research and my ideas, and for pushing me harder than anyone else. To my sister-in- law, thank you for always being there during good times and bad times; and of course for giving

V me the most amazing niece in the world. To my brother, you are the glue that holds everything together. Many times I would have crumbled without your support. Brothers for life.

To my mother, you are the guiding star I follow every day. Your light shines as bright today as it did when I was a child. It is with your love and wisdom that I am who I am today. To you I owe this manuscript. I promise you that I will always aspire for greatness, with my family and my life. I love you.

Preface

This thesis has been organized as a series of manuscripts (see the list below) which have been published in peer-reviewed scientific journals. As identified by primary authorship, all manuscripts were written by Ethan Tumarkin with critical comments and revision by Eugenia Kumacheva and corresponding collaborators. The contributions of other authors are provided in detail below.

Chapter 1 Microfluidic Studies of Biological Phenomena

The results in this chapter are mainly from manuscripts published in 1Chemical Society Reviews, 2009, 8, 2161, and 2Small, 2012, In Press

Authors: 1Ethan Tumarkin, and Eugenia Kumacheva

2Diego Velasco, Ethan Tumarkin, and Eugenia Kumacheva

Contributions: 1E. Tumarkin contributed to the article writing and figure design and preparation.

2E. Tumarkin contributed to the article writing and figure design and preparation. D. Velasco wrote the manuscript and helped with figure preparation.

Chapter 3 High-Throughput Combinatorial Cell Co-Culture Using Microfluidics

The results in this chapter are mainly from manuscripts published in Integrative Biology, 2011, 3, 653.

Authors: Ethan Tumarkin, Lsan Tzadu, Elizabeth Csaszar, Minseok Seo, Hong Zhang, Anna Lee, Raheem Peerani, Kelly Purpura, Peter Zandstra, and Eugenia Kumacheva

Contributions: E. Tumarkin contributed to the paper by designing and carrying out experiments, data analysis and interpretation, and article writing. L. Tzadu, M. Seo, and H. Zhang helped with microfluidic experiments. E.

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Csaszar, R. Peerani, and K. Purpura helped with cell culture, cell analysis, and interpretation. A Lee helped with the preparation of the manuscript. P. Zandstra provided guidance and suggestions on experimental design, and article writing.

Chapter 6 Temperature-Controlled 'Breathing' of Carbon Dioxide Bubbles

The results in this chapter are mainly from manuscripts published in 1Lab on a Chip, 2011, 11, 3545 and 2Chemical Communications, 2011, 47, 12712.

Authors: 1Ethan Tumarkin,n, Zhihong Nie, Jai Il Park, Milad Abolhasani, Jesse Greener, Barbara Sherwood Lollar, Axel Guenther, and Eugenia Kumacheva

2Ethan Tumarkin, Zhihong Nie, Jai Il Park, and Eugenia Kumacheva

Contributions: 1E. Tumarkin contributed to the paper by designing and carrying out experiments, data analysis and interpretation, and article writing. Z. Nie and J. Park help with experimental design and microfluidic experiments. M. Abolhasani assisted with simulation design and analysis. J. Greener performed ATR-FTIR experiments. B. Sherwood Lollar and A. Guenther provided guidance and suggestions on experimental design, data interpretation, and article writing.

2E. Tumarkin contributed to the paper by designing and carrying out experiments, data analysis and interpretation, and article writing. Z. Nie and J. Park help with experimental design and microfluidic experiments.

Chapter 7 Development and Applications of Microfluidic Reactors with Multiple Reconfigurable Analytical Probes

The results in this chapter are mainly from manuscripts published in 1Analyst, 2012, 137, 444 and 2Lab on a Chip, 2012, 12, 696.

Authors:

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1Jesse Greener*, Ethan Tumarkin*, Michael Debono, Chi-Hang Kwan, Milad Abolhasani, Axel Guenther, and Eugenia Kumacheva, Co-First Author

2Jesse Greener, Ethan Tumarkin, Michael Debono, A. Dicks, and Eugenia Kumacheva

Contributions:

1E. Tumarkin contributed to the paper by designing and carrying out experiments, data analysis and interpretation, and article writing. J. Greener contributed to the paper by designing and carrying out experiments, data analysis and interpretation, and article writing. M. Debono assisted with experiments. C.H. Kwan and M. Abolhasani assisted with simulation design and analysis. A. Guenther provided helpful guidance on experimental design and, as well as assisted with data interpretation and article writing.

2E. Tumarkin contributed to the paper by designing experiments, data analysis and interpretation, and supervision of M. Debono (undergraduate student). J. Greener contributed to the paper by designing experiments, data analysis and interpretation, and article writing. M. Debono performed the experiments. A. Dicks provided valuable guidance on manuscript preparation and article writing.

Publications during Ph.D. Studies

Peer Review Publications

1. D. Velasco, E. Tumarkin, E. Kumacheva, Microfluidic Encapsulation of Cells in Polymer Microgels, Small, 2012, In Press

2. E. Tumarkin, L. Tzadu, E. Csaszar, M. Seo, H. Zhang, A. Lee, R. Peerani, K. Purpura, P. Zandstra, E. Kumacheva, High-Throughput Combinatorial Cell Co-Culture Using Microfluidics, Integrative Biology, 3, 653-662, (2011), Cover

3. Greener, E. Tumarkin, M. Debono, E. Kumacheva, A Microfluidic Platform for University- Level Analytical Chemistry Laboratories, Lab on a Chip, 12, 696-701 (2012)

4. Greener*, E. Tumarkin*, M. Abolhasani, A. Guenther, E. Kumacheva, Development and Applications of a Microfluidic Reactor with Multiple Reconfigurable Analytical Probes, Analyst, 137, 444-450 (2012), Co-First Author

5. E. Tumarkin, Z.H. Nie, J.I. Park, E. Kumacheva, Temperature Mediated Generation of Armoured Bubbles, Chemical Communications, 47, 12712-12714, (2011)

6. E. Tumarkin, Z.H. Nie, J.I. Park, M. Abolhasani, J. Greener, B. Sherwood-Lollar, A. Guenther, E. Kumacheva, Temperature-Controlled 'Breathing' of Carbon Dioxide Bubbles, Lab on a Chip,11, 3545-3550, (2011)

7. A. Lee, G.F.S. Andrade, A. Ahmed, M.L. Souza, N. Coombs, E. Tumarkin, K. Liu, R. Gordon, A.G. Brolo, E. Kumacheva, Probing Dynamic Generation of Hot-Spots in Self- Assembled Chains of Gold Nanorods by SERS, Journal of the American Chemical Society,133, 7563-7570, (2011)

8. A. Lee, S. Dubinsky, E. Tumarkin, M. Moulin, A.A. Beharry, E. Kumacheva, Multifunctional Hybrid Polymer-Based Porous Materials, Advanced Functional Materials, 21, 1959–1969, (2011), Feature Article

9. A. Kumachev, J. Greener, E. Tumarkin, E. Eisner, P. W. Zandstra, E. Kumacheva, High- throughput Generation of Hydrogel Microbeads with Varying Elasticity for Cell Encapsulation, Biomaterials, 32, 1477-1483, (2011)

10. N. Raz, J. Li, L. Fiddes, E. Tumarkin, G. Walker, E. Kumacheva, Microgels with an Interpenetrating Network Structure as a Model System for Cell Studies, Macromolecules, 43, 7277-7281, (2010)

11. L. Fiddes, N. Raz, S. Srigunapalan, E. Tumarkin, V. Parkharenko, C. Simmons, A. Wheeler, E. Kumacheva, A Circular Cross-Section PDMS Microfluidics System for Replication of Cardiovascular Flow Conditions, Biomaterials, 13, 3459-3464, (2010)

12. J. Park, E. Tumarkin, E. Kumacheva, Small, Stable, and Monodispersed Bubbles Encapsulated with Biopolymers, Macromolecular Rapid Communications, 31, 222–227, (2010)

13. E. Tumarkin, E. Kumacheva, Microfluidic Generation of Microgels from Synthetic and Natural Polymers, Chemical Society Reviews, 38, 2161 – 2168, (2009), Cover

Table of Contents

Acknowledgments...... IV

Preface...... VII

Table of Contents ...... XII

List of Tables ...... XVI

List of Figures ...... XVII

Chapter 1 Microfluidic Studies of Biological Phenomena ...... 1

1.1 Why Encapsulate? ...... 1

1.2 Two-dimensional vs. Three-dimensional Cell Culture ...... 2

1.3 Development of Artificial Microenvironments for Controlling Cell Behaviour ...... 5

1.4 Microfluidic Generation of Cell-Laden Microenvironments ...... 6

1.4.1 Routes to Microfluidic Emulsification...... 7

1.4.2 Microfluidic Generation of Cell-Laden Microgels ...... 11

1.4.3 Microfluidic Emulsification of Cell Suspensions ...... 13

1.4.4 Encapsulation of Cells in Microgels ...... 17

1.4.5 Applications of Cell-Laden Microgels...... 26

1.4.6 Future work in Microfluidic Encapsulation of Cells in Microgels ...... 28

Chapter 2 Materials and Methods ...... 39

2.1 Materials ...... 39

2.1.1 Biopolymers ...... 39

2.1.2 Gases ...... 39

2.1.3 Low Molecular Weight Compounds ...... 40

2.1.4 Synthetic Particles ...... 40

2.1.5 Microfabrication ...... 40

2.2 Methods ...... 40

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2.2.1 Mask Design ...... 40

2.2.2 Microfabrication of Negative Masters ...... 41

2.2.3 Fabrication of Microfluidic Devices ...... 42

2.2.4 Microfluidic Experiments ...... 43

2.3 Characterization ...... 47

2.3.1 Optical Microscopy Imaging ...... 47

2.3.2 Size Distribution of Bubbles and Droplets ...... 47

2.3.3 Characterization of Cell-Laden Agarose Microgels ...... 47

2.3.4 Flow Cytometry ...... 48

2.3.5 Characterization of Agarose Solutions ...... 51

2.3.6 On-Chip Characterization of Chemical Reactions ...... 52

Chapter 3 ...... 55

3.1 Introduction ...... 55

3.2 Results and Discussion ...... 58

3.3 Controlling Encapsulation Efficiency ...... 60

3.4 Co-encapsulation of MBA2 Cells and M07e Cells ...... 68

3.5 Co-encapsulation of Heterogeneous Population of Primary Human Hematopoietic Cells with MBA2 Cells ...... 72

3.6 Pre-Microfluidic Experiment Considerations ...... 75

3.6.1 Wetting of the Microfluidic Channel ...... 75

3.6.2 Maintaining Cell Viability ...... 76

3.6.3 Bimodal Distribution of Particle Size ...... 77

3.7 Conclusions...... 78

Chapter 4 ...... 83

4.1 Introduction ...... 83

4.2 Results and Discussion ...... 87

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4.2.1 Characterization of Macroscopic Agarose Gels ...... 87

4.3 Microfluidic Encapsulation of AML2 Cells ...... 97

4.4 Conclusions...... 102

Chapter 5 ...... 108

5.1 Characterization of On-Chip Reactions and Integrated Analytical Tools ...... 108

5.2 Microfluidic Studies of Carbon Dioxide Dissolution ...... 109

5.3 Summary ...... 110

Chapter 6 ...... 114

6.1 Introduction ...... 114

6.2 Background ...... 118

6.3 Experimental Design ...... 119

6.4 Results and Discussion ...... 123

6.4.1 Bubble Breathing in Water ...... 123

6.4.2 Bubble Breathing in Salt Water and Ocean Water ...... 125

6.4.3 Bubble Breathing in Dimethyl Ether of Poly(ethylene glycol) ...... 128

6.4.4 Generation of Armoured Bubbles ...... 129

6.5 Conclusions...... 136

Chapter 7 ...... 142

7.1 Introduction ...... 142

7.2 Experimental Design ...... 145

7.2.1 Integration of Probes with the MF Reactor...... 145

7.2.2 Validation of Measurements of pH ...... 146

7.2.3 Validation of FT-IR characterization ...... 148

7.2.4 Validation of Temperature Measurements...... 150

7.3 Applications of the MF Reactor Integrated with Multiple Probes ...... 154

7.3.1 Titration Reactions ...... 154 XIV

7.3.2 Titration in a Microfluidic Format ...... 154

7.3.3 Titration of a Strong Acid with a Strong Base ...... 156

7.3.4 Titration of Weak Acid with Strong Base...... 159

7.3.5 Titration of a Diprotic Acid ...... 162

7.3.6 Titration of a Strong Acid with a Strong Base with Simultaneous On-Chip Temperature Measurements ...... 164

7.4 Temperature-Dependent Change in the pH of Tris Buffer ...... 165

7.5 pH-Dependent Variation in the Intensity of IR Signals ...... 167

7.6 Reaction of Carbon Dioxide with Water in the Presence of Tris ...... 167

7.7 Conclusions...... 170

Chapter 8 ...... 174

8.1 Summary and Conclusions ...... 174

8.2 Future Work ...... 176

List of Tables

Table 3.1 Calculation of the fraction of droplets, P(χ), expected to contain χ cells, for a cell concentration of 2x106 cells/mL at varying droplet diameters. λ is the average number of cells per droplet...... 64

Table 3.2 Calculation of the fraction of droplets, P(χ), expected to contain χ cells, for a cell concentration of 8x106 cells/mL, at varying droplet diameters. λ is the average number of cells per droplet...... 65

Table 7.1 Summary of the Titration of a strong acid (HCl) and a strong base (NaOH) ...... 158

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List of Figures

Figure 1.1 Schematic drawings of various types of microfluidic droplet generators the (a) flow- focusing, (b) T-junction, (c) terrace-like, and (d) and co-flowing stream geometries. The droplet and the disperse phases are labeled as A and B, respectively. Adapted with permission from reference 56. Copyright © 2009 Royal Society of Chemistry...... 9

Figure 1.2 (a) Optical microcopy images of the nodules on the thread of gelled sodium alginate (b) and the droplets of non-gelled sodium alginate. The continuous phase in (a) is mineral oil and in (b) it is undecanol. Scale bar is 300 μm. Adapted with permission from reference 56. Copyright © 2006 American Chemical Society...... 10

Figure 1.3 (a) A schematic of the MF device utilized in the encapsulation of Jurkat cells. A suspension of cells in cell media was introduced into the inlet labeled ‘cells’. The cell suspension was mixed with a second stream of sterile cell media (labeled ‘media’) for on-chip dilution of the cells. The two streams were then broken into droplets at the orifice due to shear forces imposed by the continuous phase (fluorinated oil). (b) Optical microscopy image of the generation of the cell laden droplets. Scale bar is 100 µm. Adapted with permission from reference 64. Copyright © 2008 Elsevier...... 14

Figure 1.4 (a) A highly ordered pattern of particles, which is formed in a MF channel due to hydrodynamic interactions imposed by fluidic streamlines. The ordered pattern was demonstrated for particles (top) and Jurkat cells (bottom). Both the particles and the cells self- organized in the microchannel into a diagonal/alternating pattern. (b) The distance between the particles and cells corresponded to a droplet volume of 14.6 pL. (c) A graph representing the fraction of droplets containing a specific number of particles. Adapted with permission from reference 70. Copyright © 2008 Royal Society of Chemistry...... 17

XVII dead cells) and propidium iodide (red dye, staining dead cells only) and then observed using bright field and confocal fluorescence microscopy. Yeast cell-laden microgels gelled overnight at 30 oC (a) and after treatment with 70% ethanol to kill all encapsulated cells prior to staining (b). Green staining corresponds to living cells; red or yellow staining marks dead cells. Adapted with permission from reference 77. Copyright © 2011 Elsevier...... 19

Figure 1.7 Coalescence induced gelation of Ca2+-Alginate microgels. (a) Schematic of the micro nozzle array used in the generation of droplets containing 1.5% sodium alginate and Jurkat cells in HEPES buffer saline, and droplets containing 0.1M CaCl2. Gelation was induced upon the coalescence of the two types of droplets. (b) and (c) represent optical microscope images of cell- free microgels and cell-laden alginate microgels, respectively. Adapted with permission from reference 82. Copyright © 2005 Elsevier...... 22

Figure 1.8 Cell-laden agarose microgels. (a) Fluorescence optical microscopy image of 100 m- diameter agarose microgels encapsulating mES cells. Prior to injection into the MF device, two populations of mES cells were fluorescently labeled green and red, respectively. Fluorescent labeling was done to compare the encapsulation ratio of the two populations at varying flow rate ratios of the two cell-laden agarose streams. (b) Optical microscopy image of embryoid bodies formed from mES cells encapsulated in agarose microgels after 4.5 days after encapsulation. Scale bar is 100 µm. Adapted with permission from reference 19. Copyright © 2011 Royal Chemical Society...... 23

Figure 1.9 Optical microscope images of cell-laden gelatin-Ph core-shell microgels. (a) Encapsulation of cell-laden unmodified gelatin microgels in modified gelatin-Ph microgels. (b) 1 day of post-encapsulation, and (c) 4 days. (d) 1 day after seeding L929 cells on the surface of the gelatin-Ph microgels. Adapted with permission from reference 20. Copyright © 2011 American Institute of Physics...... 25

and the ratio of the respective flow rates, Q1 and Q2, was changed in a throughput manner. (c) Two agarose solutions, one containing a suspension of cells, and the other containing a bioactive reagent are supplied in the MF device at varying flow rates Q1 and Q2. Copyright © 2012 WILEY-VCH...... 28

Figure 3.1 Microfluidic encapsulation of cells in agarose microgels. (a) Optical image of the MF device used for the generation of cell-laden agarose microgels. Agarose solutions were maintained at 37 oC in a temperature controlled incubator throughout the experiment. The outlet tubing was embedded in a temperature controlled water circulator tubing at 2 oC to ensure gelation of the agarose droplets prior to collection in HBSS buffer. (b) Schematic of MF device shown in (a). A serpentine downstream channel with a length of 250 mm is not shown in the figure. (c) The distribution of sizes of cell-laden droplets of 2 wt. % agarose solution (dotted curve) and of the corresponding agarose microgel (solid curve). Scale bar is 4 mm...... 59

Figure 3.2 Theoretical Poisson distribution curves for the number of cells per microgel were generated by calculating the average number of cells per droplet at a constant cell concentration and droplet volume and subsequently fitting the data to a Poisson distribution by using the Poisson function avaible in Microsoft Excel 2007. Theoretical Poisson distribution of the fraction of microgels containing a particular number of cells (a) at a constant cell concentration (8 x 106 cell/mL) and a variable droplet diameter and (b) at constant droplet diameter of 110 µm and variable cell concentration...... 61

Figure 3.3 Encapsulation efficiency of the microfluidic encapsulation of cells at varying cell concentration and droplet diameter. Experimental (blue) and theoretical (grey) fractions of droplets encapsulating at least one cell are plotted for 70 and 110 µm-diameter droplets at cell concentrations of 2x106 and 8x106 cells/mL. The theoretical fraction of cell-laden droplets was calculated using eq. 2...... 62

Figure 3.4 Comparison of the number of encapsulated cells per droplet compared to theoretically expected Poisson distribution. The graph shows the number of cells per microgel determined experimentally (filled symbols) and theoretically using the Poisson distribution (empty symbols). The concentration of cells in the feed suspension was 4x106 (dash lines) and 8x106 cells/mL (solid lines). Each experimentally acquired data set was compiled from approximately 300 microgels...... 64 XIX

Figure 3.5 Cell-laden agarose microgels. (a) Fluorescence optical microscopy image of mSC cell-laden 100 m diameter agarose microgels. Encapsulation occurred at a total cell concentration of 8x106 cells/mL in the droplet phase. The cells appearing to reside outside the microgel are out of the focus plane of the microscope. (b) Optical microscopy image of embryoid bodies formed from mES cells encapsulated in agarose microgels after 4.5 days after encapsulation. The flow rate of the oil and aqueous phases were 1.2 and 0.09 mL/hr, respectively. Scale bar is 100 µm...... 65

Figure 3.6 Effect of dilution on encapsulation of individual cell populations. Mixing of two streams of distinct cell suspensions results in the dilution of individual cell populations. For example, for two suspensions, each containing 8 x 106 cells/mL, mixing of streams supplied in the MF device at equal flow rates (QG:QR =1:1) reduced the concentration of individual cell populations to 4 x 106 cells/mL. The graph shows good agreement between the theoretical Poisson distribution for the percentage of 110 µm-diameter cell-laden microgels at a cell concentration of 4 x 106 cells/mL and the experimental results obtained for the encapsulation of green and red cells in 110 µm-diameter microgels at the flow rate ratio of the corresponding suspensions of 1:1...... 66

4:1, (e) QR:QG = 1:0. Gating was determined by positive controls comprising cells labelled with only one dye (R:G = 1:0 and 0:1). (F) Fraction, α, of green and red cells encapsulated at different ratios of QR:QG. Light and dark green bars show the fraction of encapsulated green cells, determined by image analysis and flow cytometry, respectively. Light and dark red bars represent the fraction of encapsulated red cells, determined by image analysis and flow cytometry, respectively. Fluorescence intensity scale is defined by the sorting parameters. Scale bar is 100 µm...... 68

Figure 3.8 Variation in the viability of M07e human megakaryoblastic leukemia cells examined at varying encapsulation ratio of MBA2 to M07e cells. (a) Schematic of co-culture of MBA2 and M07e cells in different number ratios. MBA2 cells secrete IL-3 which is required for the survival of both cell populations. Increase in the relative number of MBA2 cells results in an increase in M07e survival rate, due to the local increase in IL-3. (b) Viability of M07e cells plotted as a function of the number ratio of MBA2 cells-to-M07e cells encapsulated in 100 m-diameter agarose microgels and analyzed by flow cytometry. Asterisk denotes non-encapsulated M07e cells in the presence of exogenous IL-3. (c) Representative flow cytometry histograms of cellular uptake of 7-AAD fluorescence intensity for varying M07e co-culture conditions 4.5 days after encapsulation. X-axis represents fluorescence intensity of cells stained with 7-AAD. Numerical percentages represent the mean and standard error of M07e cellular uptake of 7-AAD dye from at least three independent experiments...... 71

Figure 3.9 Viability of encapsulated M07e cells at varying media volume. Microgels containing only MBA2 cells and only M07e cells mixed in a 1:1 ratio at varying media volume. Asterisk denotes non-encapsulated M07e cells in the presence of exogenous IL-3...... 72

Figure 3.10 Co-encapsulation of MBA2 cells with UCB cells were used to determine the survival effect of IL-3 on sub-populations within a hematopoietic culture system. (a) Schematic of co- culture of MBA2 cells and UCB cells. (i) Encapsulation of solely UCB cells without exogenous IL-3 results in poor cell rescue. (ii) Addition of exogenous IL-3 (10 ng/mL) results in low rescue of several UCB phenotypes. (iii) Co-encapsulation of MBA2 cells and UCB cells results in low, moderate, and high rescue of specific UCB phenotypes. (b) % Rescue of specific lineages of hematopoietic UCB cells co-encapsulated with MBA2 cells. % rescue of specific UCB phenotypes were measured 3 days after encapsulation in a non-supportive media using eq. 1. This is a representative experiment conducted from at least two independent experiments (n = 2)...... 75

Figure 3.11 Optical microscopy image of a bi-modal population of microgels containing two distinct sizes. Scale bar is 100 µm...... 77

Figure 4.1 Variation in the viscosity of agarose solution with increasing concentration of the biopolymer in PBS solution. Measurements were conducted at 37 oC. 1, 2, 3, wt. % solutions

XXI were measured with a spindle spin rate of 60 RPM, 4 wt. % was measured at a spindle spin rate of 20 RPM, and 5 wt. % was measured at a spindle spin rate of 8 RPM...... 89

Figure 4.2 Variation in the Young’s modulus of agarose microgels at r.t. (●) and 37 oC (▲), and macroscopic agarose gels at r.t. (■) at increasing concentration of the biopolymer in PBS solution. In some instances error bars are smaller than the data points...... 91

Figure 4.3 Optical microscopy images of cell laden agarose gels. Day 0: Images of single cells embedded inside agarose gels at varying Cag. Images were recorded directly after gelation of the agarose. Day 7: Optical microscopy images of cell-laden agarose gels on Day 7 at varying Cag. Scale bar is 100 µm...... 92

Figure 4.4 Fluorescent images of cells encapsulated in 1 wt. % (top) and 3 wt. % (bottom) agarose on Day 7 of cell culture. (a,g) Ethidium homodimer-1 was used to generate red fluorescence in non-viable cells. (b,h) Viable cells labelled with Calcein AM (green). (c,i). Hoescht dye was added to all samples to stain DNA of both viable and non-viable dyes. (d) Bright field image of the cell colony shown in a-c, d-e. (e) Composite image overlaying fluorescence of both viable (Calcein AM) and dead (Ethidium homodimer-1) cells shown a and b (f) Composite image overlaying fluorescence of Hoescht stained cells and dead (Ethidium homodimer-1) cells appearing in a and c. (k) Composite image overlaying fluorescence of both live (Calcein AM) and dead (Ethidium homodimer-1) cells appearing g and h. (l) Composite image overlaying fluorescence of hoescht stained cells and dead (Ethidium homodimer-1) cells shown in g and i. Scale bar is 50 µm...... 94

Figure 4.5 Structure of WST and its reduction to a Formazan Dye...... 96

Figure 4.6 Graph of the absorbance of Formazan Dye at varying agarose concentration. Metabolically active cells convert WST to Formazan Dye. An increase in the absorbance at 450 nm is due to an increase in the number of viable cells. 0 wt. % agarose represents a positive control of AML2 cells suspended in agarose-free cell media. Absorbance was normalized with respect to the average absorbance of Formazan Dye in 5 wt. % agarose...... 97

Figure 4.7 Schematic of the MF device for the generation of cell-laden agarose microgels with tunable elasticity. (a) MF device comprising one inlet channel for the disperse phase. (b) MF device comprising two inlet channels for injection of the disperse phase. The height of both XXII devices was 150 µm. The width of the horizontal channels supplying the mineral oil phase and the width of the channel at the point of the T-junction was 150 and 20 µm, respectively. The length of the mixing channel prior to the T-junction was 250 mm...... 99

Approximately 300 microgels (formed at Cag = 2 wt. %) were analyzed to determine the encapsulation efficiency. The diameter of the precursor droplets was 110 +/- 2.8µm. QOil = 1.2 mL/hr, Qag = 0.1 mL/hr...... 101

Figure 4.9 Optical microscopy images of 1 and 2 wt. % agarose microgels encapsulating AML2 cells in cell culture media on Day 0 and 7. The concentration of cells in the feed suspension was 1x106 cells/mL. The scale bar is 100 m...... 102

Figure 6.1 Temperature-dependent shrinkage and expansion of CO2 bubbles in liquid. Decrease in temperature leads to increase in the solubility of CO2 in the liquid and reduction in bubble size. With increase in temperature, the solubility of CO2 n the liquid decreases, leading to bubble expansion...... 119

Figure 6.2 (a) Schematic of the MF device. Regions 1 and 3 are cooled by placing underneath them a hollow copper block. Region 2 is heated by placing underneath it a heating module. (b) Variation in temperature of the continuous aqueous flowing through the MF device, plotted as a function of the distance from the orifice. Open squares represent experimentally determined temperature. The dotted line shows simulated temperatures at varying distance from the orifice...... 121

Figure 6.3 Schematic of the accessories for the temperature control in the MF device. The position of the heating module and the cooling module are placed below the MF device fabricated in PDMS: (a) Side view and (b) perspective view of the setup. The cooling module is inserted underneath Regions 1 and 3, and the orifice, the heating module is applied to Region 2. The temperature of 0 oC of the cooling module is controlled by flowing through it a mixture of

XXIII glycerol and water (1:4 v/v) using a temperature-controlled water circulator. The temperature of the heating module of 35 oC is controlled by a temperature controller...... 122

Figure 6.4 Optical microscopy images of “breathing” CO2 bubbles. The images were acquired (a) immediately after the orifice; (b) 125 mm away from the orifice in the cooled Region 1; (c) 235 mm away from the orifice in the heated Region 2; (d) 447 mm away from the orifice in the cooled Region 3. PCO2= 48.3 kPa, QC = 2.5 mL/hr. The scale bar is 100 µm...... 123

Figure 6.5 (a) Bubble volume plotted as a function of the distance from the orifice. The lines are given for eye guidance. (b) Volume of bubbles plotted as a function of the temperature of the continuous phase. Numbers represent points shown in (a). The graph demonstrates a contraction- expansion-contraction cycle for the change in bubble volume. (c) Variation in CO2 bubble volume at 23 oC, plotted as a function of the distance from the orifice. The bubbles were generated at a PCO2 = 48.3 kPa and QC = 2.5 mL/hr...... 125

Figure 6.6 (a) Variation in bubble volume in ocean water () and an aqueous 0.7M NaCl solution (). The lines are given for eye guidance. (b) Measurement of the pH of the continuous phase were taken at Points 1, 2, and 3 (as in (a)), corresponding to temperatures 1, 31, and 3oC, respectively. The bubbles were generated at PCO2 = 48.3 kPa, and QC = 2.5 mL/hr...... 127

Figure 6.7 (a) Variation in the volume of CO2 bubbles in DEPG and in the temperature of DEPG, plotted as a function of the distance from the orifice of the MF device. The lines are given for eye guidance. Images (from left to right) show bubbles at temperatures of 1, 31, and 3 oC, respectively. (b) Variation in the amount of dissolved CO2, plotted as a function of the experimentally determined (black symbols) and simulated (white symbols) temperatures. (c) Variation in bubble volume in DEPG, plotted as a function of the distance from the orifice at 23 o C. PCO2 = 82.7 kPa, Qc = 2.5 mL/hr. The scale bar is 50 µm...... 129

Figure 6.8 (a) Optical microscopy images of CO2 bubbles flowing through the microchannels through zones at varying temperatures. (b) The variation in temperature in the microchannel (top) and corresponding variation in the average volume of bubbles (bottom), both plotted as a function of the distance from the orifice of the MF device: (■), (●), (▲) correspond to three sets of experiments operated at different temperature profiles. The vertical dashed lines indicate the

XXIV

boundaries between the three zones with different temperatures. PCO2 = 20.7 kPa. The flow rate of the continuous aqueous phase, QL is 3.0 mL/hr...... 132

Figure 6.9 (a) Optical microscopy images of CO2 bubbles armoured with a shell of PS-co-PAA particles. Inset shows a high magnification image of an armoured bubble. (b) Size distribution of armoured bubbles. The armoured bubbles were obtained at the temperature of the liquid in the o microchannel of T = 28 C, PCO2 = 79.3 kPa, QL = 24 mL/hr, pH = 14, Cp = 1.2 wt. %. Scale bar is 100 μm (a) and 30 μm (inset)...... 134

Figure 6.10 The relative change in the volume of CO2 bubbles (a) and the diameter of armoured bubbles (b) as a function of temperature. Insets in (b) are typical optical microscopy images of armoured bubbles obtained at corresponding temperature (indicate by arrows). PCO2 = 79.3 kPa,

QL = 24 mL/hr, pH = 14, Cp = 1.2 wt. %...... 134

Figure 7.1 (a) Schematic of the MF reactor. The position of the three probes is marked with open circles at points P1, P2, and P3. P1, P2, and P3 at a distance of 49, 277, and 291 mm, respectively, from the inlet of the MF reactor. (b) A schematic of the cross-section of the MF reactor interfaced with a (i) temperature probe, (ii) pH probe, (iii) ATR-crystal. The ATR crystal is interfaced with the bottom of the microchannel. (c) Optical microscopy image of the fragment of the microchannel interfaced with a circular ATR crystal at P3. Arrows indicate the direction of flow. Scale bar is 500 μm. (d) A photograph of a MF reactor with three integrated probes. Only one inlet is connected to avoid obstruction of the device...... 146

Figure 7.2 Response of the integrated pH probe to (a) the variation in the pH of the liquid. The alternating pH values correspond to the alternating introduction of solutions of HCl (pH = 2.3) and NaOH (pH = 11.5). The time between the dashed lines indicates the time required for the probe to respond to the new pH value. Arrows indicate the time of stopping of the flow of either the acidic or basic solution and the simultaneous introduction of the counterpart solution. Each solution was supplied at the flow rate Q = 2 mL/h. (b) Measurement of pH of the solution of HCl (pH = 2.3) supplied to the MF reactor at varying flow rates...... 148

XXV required for the absorbance at 1060 cm-1 to change (~ 15 s). Arrows indicate the time of stopping of the flow of either the water, or the Tris solution and the simultaneous introduction of the other liquid. Each solution was supplied at the flow rate Q = 3 mL/h (b) Variation in absorbance (1060 cm-1) vs. flow rate of the 50 mM Tris solution. Error bars were obtained by determining the standard deviations in absorbance, based on five independent measurements for each flow rate of the liquids. Spectra were collected using 8 scans at 10 kHz and 8 cm-1 spectral resolution...... 150

Figure 7.4 Measurement and control of temperature in the MF reactor. (a) Variation in temperature of the liquid flowing at a flow rate 4 mL/h by using a feedback temperature control process. The temperature of the liquid is cycled between 25 and 50 oC. (b) Temperature achieved at point P1 without feedback control, plotted as a function of the flow rate of the liquid. Filled circles and a dotted line show experimental results and the results of simulations, respectively. Squares show the temperature at Point 1 achieved with feedback temperature control. (c) An exemplary simulation result of heat transfer from a heating pad placed below the 1 mm MF device. The flow rate of the liquid (water) is 0.9 mL/h, the heating pad was set to 50 oC. Inlets and probe positions are shown on the image. The inset shows an enlarged image of the temperature of the flowing liquid at in the inlet. Scale bar is 4 mm...... 152

Figure 7.5 Microfluidic titration of a strong acid with a strong base. Titration curves for [NaOH]i=0.05M, and [HCl]i equal to (a) 0.025M (■), (b) 0.035M (▲) and (c) 0.055M (●). The total volumetric flow rate was QT = 2.0 mL h-1. The dashed lines show theoretical curves calculated using eqs. 6 and 7...... 157

Figure 7.6 Microfluidic titration of a weak acid with a strong base. (a) Titration curve for

[CH3COOH]=1M, [KOH] =1M. Data acquisition was conducted 2-3 min after changing the flow rates of the liquids. The pKa is measured from the first inflection point (where the change in pH is minimum) and the pHe is measured from the second inflection point (where the change in pH is maximum)...... 161

Figure 7.7 Microfluidic titration of a strong base with a multiprotoic acid. (a) Titration curve for

[CH2(COOH)2] = 0.09M, [KOH] = 1.00M. The data were acquired based on the two independent experiments. Acquisition was conducted 2-3 min after changing the flow rates of the liquids. The values of QKOH/QT yielding pH = pKa1 and pH = pKa2 were determined by eye at the inflection XXVI

points in on the titration curve, where the slope was lowest. The values of QKOH/QT corresponding to pH = pHe1 and pH = pHe2 were determined by eye at the inflection points where the slope was largest. (b) The exact QKOH/QT values yielding pH = pKa1 and pH = pKa2 were determined by the local minima in the first derivative plot of (a). The exact QKOH/QT values leading pH = pHe1 and pH = pHe2 are determined by local maxima in the first derivative plot of (a)...... 163

Figure 7.8 Variation in temperature (●) and pH (■) measured for mixing aqueous solutions of HCl and NaOH. The ratio of base/acid concentrations was controlled by varying the flow rate ratio of the 0.25 M solution of NaOH and 0.5 M solution of HCl from 0:1 to 1:0. The total volumetric flow rate of the liquid was 4 mL/h. The pH titration curve shows an equivalence point at the [HCl]:[NaOH] molar ratio of 1:1...... 165

Figure 7.9 (a) Variation in pH of Tris buffer solutions with varying temperature. Buffer solutions pH values of 7.1 (◊) and 7.9 (■) at room temperature. The flow rate of each buffer solution was 0.5 mL/h. (b) FTIR spectra of 200 mM Tris buffer solution after diluting it with 1M HCl (aq). The concentrations ratio of [Tris]:[HCl] (mM) were (1) 200:0, (2) 195:24, (3) 186:70, (4) 167:167, (5) 148:259, (6) 121: 394, (7) 113:437, (8) 100:500, (9) 99:506, (10) 98:512. These mixing ratios resulted in the corresponding measured pH values of (1) 11.2, (2) 10.1, (3) 9.4, (4) 8.9, (5) 8.6, (6) 8.1, (7) 7.8, (8) 7.1, (9) 2.1, (10) 1.6. Arrows show a change in absorbance with a reduction in pH. (c) Absorbance of peaks shown in (b) vs. measured pH. Error bars in (a) and (c) were determined from the standard deviation from 3 separate measurements, but are not shown because they were smaller than the data points...... 166

XXVII

-1 spectrum of CO2 (aq) in Tris buffer (pH = 6.4). The peak at 2342 cm (indicated with an arrow) corresponds to the asymmetric vibration of O-C-O. The spectra shown in (d) and (e) were acquired from a single measurement comprised of 16 scans at 10 kHz and spectral resolution 4 cm-1. The window region of low transmission of the diamond ATR (2200 to 1800 cm-1) has been excluded from both (d) and (e)...... 169

XXVIII

Chapter 1

Microfluidic Studies of Biological Phenomena

1.1 Why Encapsulate?

The encapsulation of cells in micrometer-size three-dimensional hydrogels offers the ability to control the shear forces imposed on cells, the ease of cell visualization, the reconfigurability of the cell-laden hydrogel modules, and the transport of oxygen, nutrients, growth factors and waste products.1 These advantageous characteristics have led to a large number of biomedical applications of microscopic cell-laden hydrogels, including clinical diagnostics, pharmaceutical research, and regenerative medicine. The transplantation of cells compartmentalized within hydrogel microcapsules was utilized as early as in 1964, in order to suppress immune rejection of transplanted cells.2 The encapsulation prevented the entry of antibodies and immune cells (T cells) into the capsule, but permitted the transfer of therapeutic products from the cells to the surrounding environment. Since then, the implantation of hydrogel beads carrying different types of cells was used for in situ production and delivery of important biological molecules, in order to treat diseases or to repair damaged tissue.3-4

Cell-laden microscale hydrogel “modules” have promising applications in tissue engineering.

Modules carrying different types of cells can be combined or reconfigured to mimic various types of tissue. Within these modules, cells show proliferation to densities of 108-109 cells cm-3, which are comparable to those of native tissue.5

The encapsulation of cells in hydrogel modules or microbeads (microgels) can also be utilized for exploratory purposes. In natural microenvironments, all cells, except those in circulation, 1

require anchorage to a matrix. Cell fate is influenced by multiple cues, including exposure to soluble growth factors, interactions with neighboring cells, and mechanical stimuli from the surrounding extracellular matrix (ECM).6 Understanding and reproduction of the properties of cellular microenvironments is vital in tissue engineering, wound healing and treatment of diseases. For instance, the malignant transformation and migration of cancer cells are strongly affected by the stiffness, porosity and architecture of the surrounding environment.7 The viability, renewal and differentiation of stem cells towards a particular lineage are dependent on the properties of cellular microenvironment (“niche”), which determine concentration gradients of signaling factors, matrix elasticity, and appropriate oxygen levels.8-10 Encapsulation and subsequent cell culture in microscopic hydrogel beads (microgels) can be used to study the role of properties of microenvironments in cell fate, in order to understand and develop an in vitro cell culture conditions that mimic specific in vivo environments. To address this goal, a high- throughput combinatorial approach to the generation of a large number of cell-laden microgel particles with varying properties is highly desirable.

1.2 Two-dimensional vs. Three-dimensional Cell Culture

Conventional methods for culturing cells in-vitro involve the formation of a two-dimensional

(2D) monolayer of cells on the surface of a plastic culture dish. Recently, the scientific discussion has intensified on the extent to which studies of cells cultured on 2D substrates can be related to in-vivo cell behavior. A growing body of evidence suggests that three-dimensional

(3D) culture environments more accurately mimic the natural environment of cells in terms of microenvironment architecture, and chemical and mechanical signal transduction.11-14 Two- dimensional culture systems suffer from several important drawbacks. In their natural environment, the majority of cells naturally reside inside complex 3D tissue matrices, which

impose chemical and mechanical cues from all directions. Culturing cells on 2D surfaces favors cell-substrate interactions on only one side of the cell, thereby imposing anisotropy in the forces felt by the cell. Consequently, cells cultured on 2D substrates show dramatically different motility when compared with studies in 3D environments.7 As a result, the encapsulation of cells in 3D environments has become an increasingly popular approach in both stem cell15 and cancer cell biology.16-17 Various biological hydrogel materials have been developed for the generation of 3D microenvironments, including agarose,18,19 gelatin,20 alginate,21 as well as naturally

22,23 ,24 derived extracellular matrix (ECM) MatrigelTM, collagen,1 and other ECM derived polymers.25 An alternative approach to 3D cultures is the generation of cellular spheroids (multi- cell aggregates) grown in suspension.26,27

One of the first realizations that 3D cell cultures provides more natural growth conditions compared with 2D monolayers was reported by Sutherland et al. in 1979.28 This group has shown that tumor cells grown into multicellular spheroids were less susceptible to anti-tumor drugs as compared to freely floating suspension cells. The effect was attributed to the following reasons: (i) cells in the inner mass of the spheroid showed greatest drug resistance as a result of the poor penetration of the drug through the outer mass of the spheroid, (ii) it was hypothesized that the drug was metabolically altered to a less toxic form by the cells in the outer mass prior to reaching the inner mass cells, and (iii) it was realized that cells in the inner cell spheroid were more hypoxic compared to cells in the outer cell mass, which led to increased resistance to the drug, although the exact mechanism of hypoxic resistance was not determined.

In a similar study, it was shown that higher resistance to anticancer drugs was observed in cells encapsulated in collagen gels and cultured as cellular spheroids, as compared to cells grown on

2D culture.29 From an anatomical and physiological point of view, cancer cells cultured in 3D via spheroid or hydrogel cultures mimic in vivo tumors to a significantly higher extent, compared to monolayer cultures. In particular, early stage tumors show appreciable similarities to tumor cell spheroids grown in 3D culture systems. In addition, the internal morphology of cell spheroids tend to develop hollow cores, which strongly resemble the internal morphology of early tumors, possibly as a result of cell death due to limited diffusion of oxygen and nutrients.

Lastly, the proliferation of tumor cells cultured in 3D is typically slower, which resembles physiological migration more closely than that of monolayer cultures.30 With the discovery and subsequent studies of the behavior of tumor cells cultured in 3D culture, there has been a great deal of research into translating these results to a wider scope such as stem cell science and tissue engineering.

The 3D microenvironment provides both physical and chemical cues to cells as a part of the overall cellular niche. It has been previously shown that the elasticity of the cellular microenvironment can dramatically impact the differentiation of mesenchymal stem cells

(MESc).8,31,32 Cells cultured on soft substrates with a Young’s modulus of ~ 0.1-1 kPa differentiated towards the neurogenic (neuron) lineage. A matrix with a Young’s modulus ~8-17 kPa governed cells differentiation towards the myogenic (muscle) lineage. Cells seeded on a hard matrix with an elasticity of 25-40 kPa differentiated towards the osteogenic (bone) lineage. This study demonstrated the implications that mechanical properties of the substrate can have on stem cell plasticity and the possibility to control cell fate by directly varying the properties of the microenvironment.33,34

1.3 Development of Artificial Microenvironments for Controlling Cell Behaviour

A deeper understanding of the complex biophysical interactions between cells and their respective environments requires the design and synthesis of increasingly diverse 3D matrix materials.35 Recently there has been an increased emphasis on designing matrix materials which can be independently tuned in their chemical and mechanical properties in order to maximize the interrogation ability of the encapsulated cells.

One approach to controlling cell behavior includes the manipulation of local chemical cues. In

2006, Yamanaka et al. developed the first protocol to reprogram differentiated cells into induced pluripotent stem (iPS) cells, which resembled (in behavior and phenotype) embryonic stem cells.36,37 Differentiated adult fibroblasts were cultured in the presence of four specific endogenous growth factors, which then gave rise to a small fraction of iPS cells. The presence of specific chemical cues reverted the fibroblasts to a state of pluripotency.

The development of materials which can provide both chemical and mechanical cues for cell culture are highly sought after. Typically, the mechanical properties of a matrix material can be tuned by varying the dry weight of the polymer. An increase in content of dry polymer leads to an increase in the elastic modulus of the gel matrix.7,38 Similarly, chemical coupling of specific growth factors or small molecules to the hydrogel matrix allows for direct integration of chemical cues into the microenvironment.18,38 A natural progression in the development of a matrix material is a material properties that can be tuned after the material has been prepared.

For instance, recently, a synthetic hydrogel matrix composed of polyethylene glycol (PEG) precursors was functionalized with RGD peptides (cell integrin-binding motif) and matrix metalloproteinase (MMP) sensitive regions (MMPs are zinc-based proteins capable of degrading 5

extra-cellular matrix proteins and are commonly associated with tissue remodeling and cell motility).38,39 The mechanical properties of the matrix were easily tuned by varying the initial amount of polymer in the hydrogel matrix, whereas the integration of RDG peptides into the matrix allowed for chemical interaction of the cells with the microenvironment. The incorporation of regions which are degradable by MMP activity, allowed for MMP producing cells to breakdown the matrix in a manner which closely resembles in vivo behavior. The encapsulated epithelial ovarian cancer cells were studied for their ability to physically interact with the matrix via the RGD sequences, as well as their ability to migrate within the microenvironment by breaking down the hydrogel matrix using of MMPs. This study demonstrated the importance of culturing cells in a dynamic 3D environment, which could not have been done with a 2D culture.

To explore a large range of properties and compositions of cellular niches, a method for producing cell-laden microgels in a high-throughput manner would be highly desirable, as it would allow for the generation of a large library of 3D cellular microenvironments..

Furthermore, a technique capable of tuning the chemical and mechanical properties during the formation of the microenvironment would provide the ability to interrogate many variables in parallel. The following section discusses advances in the use of microfluidic (MF) platform for the high-throughput generation of cell-laden microgel microenvironments.

1.4 Microfluidic Generation of Cell-Laden Microenvironments

Recently, a microfluidic (MF) platform has offered a promising strategy for the generation of cell-laden microgels via rapid, high-throughput production of highly monodisperse cell-laden droplets, which were subsequently gelled to form microgels with tunable compositions and

mechanical properties.40-43 We note that currently both droplets and microgels generated by the

MF means are being used as cellular microenvironments; however microgels have a number of crucial advantages and a broader range of applications, as discussed in Chapter 1, Section 1.4.5.

Here we highlight the current advancements in producing cell-laden microgels using MFs. We limited this section to the multi-phase MF generation of cell-laden microgels with dimensions not exceeding several 100-200 m. It should be noted that other MF techniques, primarily digital microfluidics44 and single phase projection lithography,45 have been utilized in for the encapsulation and growth of cells in microenvironments.

1.4.1 Routes to Microfluidic Emulsification

Microfluidic generation of hydrogel particles begins with the emulsification of an aqueous solution of a monomer, an oligomer, or a polymer in an immiscible non-polar liquid.

Emulsification in MF devices can occur via different mechanisms which depend on the geometry of the MF device, the macroscopic properties of the liquids, and the flow rates of the liquids. Figure 1.1 illustrates MF emulsification methods that were most frequently utilized to produce droplets with a narrow size distribution. Figure 1.1a shows a flow- focusing MF device. An aqueous phase and a non-polar liquid (labeled as A and B, respectively) are introduced to the central and side channels, respectively. A stream of the aqueous droplet phase is focused in the narrow orifice by the shear force imposed by the continuous phase. A highly periodic break-up of the thread of the disperse phase yields droplets with narrow size distribution.46 The size of droplets is controlled by varying the ratio of flow rates of the continuous-to-droplet phases, so that for a particular geometry of MF device and the selection of liquids, with increasing ratio of flow rates, the diameter of

droplets decreases.47 Based on experimental observations, the use of liquids with higher viscosities favors the formation of larger droplets with a narrow size distribution. 48

Figure 1.1b shows a MF droplet generator with a T-junction geometry.49 As the droplet phase enters the junction, the continuous phase forms a thin film layer between the disperse phase and the walls of the device, causing an increase in pressure which “squeezes” the disperse phase to form a droplet.50 The size of plugs (droplets that span the entire width of the channel) is controlled by the ratio of flow rates of the continuous-to-droplet phases: with increasing flow rate ratio, the droplets become smaller. The size of spherical (unconfined) droplets is not directly determined by the ratio of flow rates of the liquids.51 At low Reynolds number (Re ~ 1), increasing the flow rate of the continuous phase and decreasing the flow rate of the droplet phase results in the reduction of droplet size. At Re > 10, increasing the flow rate of the droplet phase leads to an increase in the frequency of droplet formation rather than affecting the size of the droplets, and the dimensions of droplets are controlled by tuning the flow rate of the continuous phase. Figure 1.1c shows a schematic of a terrace-like design of MF devices as an alternative geometry for droplet generation.52,53 At the end of the terrace, a stream of the disperse phase flows into a well and breaks into a disc-like droplet, which rapidly transforms into a spherical droplet due to the action of interfacial tension. The droplet is forced towards the exit of the MF device by a cross-flowing continuous phase. In this method, the diameter of droplets is controlled by the dimensions of the terrace channel and the ratio of the viscosities of the disperse phase and continuous phases.52,53 For a particular combination of the continuous and droplet phases, the flow rate of the disperse phase determines the frequency of droplet formation, while maintaining a specific diameter of droplets. Beyond the maximum threshold flow rate of the droplet phase, the size of

droplets dramatically increases.

Finally, Figure 1.1d shows a droplet generator, which consists of two concentric capillaries.54

The liquid-to-be-dispersed and continuous phases are supplied into the inner and outer capillaries, respectively. Droplets are generated when a stream of the continuous phase provides a sufficient shear stress to break up the disperse phase into droplets. Similar to the flow-focusing method, the diameter of droplets in the co-flowing geometry decreases with an increasing ratio of flow rates of the continuous-to-droplet phases. The effect of the ratio of the viscosities of the continuous and the disperse phases has little effect on droplet dimensions, however, it has been shown that with decreasing interfacial tension between the continuous and the droplet phases the size of droplets increases.55

Microfluidic generation of microgels imposes several important requirements to the processes of emulsification of polymer solutions and gelation of precursor droplets. A challenge in the MF emulsification of the precursor solutions is the possibility of rapid, drastic increase in their viscosity, which occurs upon mixing with a cross-linking agent or upon cooling. For example, Figure 1.2a shows an optical microscopy image of the thread of the mixed aqueous solutions of sodium alginate and CaCl2 (an ionic cross-linking agent)..

The thread of the droplet phase formed nodules and did not break up into droplets in a broad range of flow rates of the liquids, in contrast with a solution of sodium alginate for which gelation was delayed (Figure 1.2b).56 Thus for the controlled preparation of microgels by MF methods, it is imperative to separate in time (and MF space) the emulsification and gelation stages.

On the other hand, small dimensions of MF reactors make fast gelation particularly important, so that the microgels emerging from the reactor are sufficiently strong to keep their integrity, 10

especially upon their transfer into an aqueous phase. Finally, in order to keep a narrow size distribution of the microgels, it is important to avoid collisions of gelling droplets in the microchannels.

1.4.2 Microfluidic Generation of Cell-Laden Microgels

The specific requirements to the chemical and physical properties of cell-laden microgels depend on their future application. Below we summarize the general preferred characteristics of the methods used for the preparation of cell-laden microgels.

(i) It is imperative to generate microgels with dimensions not exceeding ~200 µm, in order to allow rapid delivery of oxygen, nutrients and metabolic products to the encapsulated cells;

(ii) A narrow distribution of sizes of cell-laden microgels enables control over the average number of cells per microgel, thereby assisting in studies of cell co-culture, the effect of confinement and intercellular distance, and the delivery of functional molecules to a particular number of the encapsulated cells;

(ii) The composition of the microgel and the method of microgel preparation should not affect cell viability. Typical factors causing cell mortality include strong shear forces, ultraviolet irradiation and large temperature gradients;

(iii) The method used for cell encapsulation has to be robust, reproducible, and produce a large number of cell-laden microgels per unit time.

Conventional methods used for the encapsulation of cells in microgels include bulk emulsification,57 electrostatic dripping,58 extrusion methods,59-61 and hydrodynamic dripping.62

These methods have a number of limitations, including a broad distribution of microgel sizes

(polydispersity in the range from 10 and 50%), large microgel dimensions (up to millimeters), and high consumption of reagents.

Microfluidic generation of microgels offers the capability to address all of the requirements listed above. In MF devices, microchannel dimensions typically vary from tens to hundreds of micrometers, which enables the generation of microgels with comparable dimensions.41 The resulting particles have narrow polydispersity (under particular conditions, the coefficient of variation in droplet dimensions does not exceed 1-2%).40 Microgels can be produced with various shapes and morphologies, including a uniform morphology or a capsular (core-shell) structure.56 In capsules, the external membrane of the microgel enables protection of the encapsulated species from harmful external stimuli, while a liquid core provides a suitable cellular microenvironment.3,63

Microfluidic generation of cell-laden microgels offers the capability for high-throughput variation in their composition and/or mechanical properties by supplying multiple streams of precursor liquids at varying relative flow rates. Microfluidics provides a route to the generation of tens of thousands of cellular microenvironments (libraries of cell-laden microgels with different properties), which can be studied by optical microscopy, or high-through-put screening using flow cytometry and automated fluorescence microscopy.19

A MF platform allows for the physical and chemical isolation of droplets from the atmosphere, thereby eliminating the risk of cross-contamination from bacteria or small molecules. In addition,

MF cell encapsulation can be easily carried out in inexpensive, sterile, dust-free, and disposable devices.

1.4.3 Microfluidic Emulsification of Cell Suspensions

Microfluidic encapsulation of cells in microgels starts from the emulsification of an aqueous suspension of cells in an immiscible non-polar liquid (a continuous phase). Typical non-polar liquids used in MF cell encapsulation include different types of oils such as a mineral, vegetable oil, and fluorinated oils.19,21,64 The suspension of cells contains a monomer or a polymer, which are used for subsequent gelation of the droplets by chemical or physical means.

The encapsulation of cells in droplets can been achieved in MF droplet generators with different geometries, including a flow-focusing device, a T-junction, co-axial capillaries and a micro- nozzle cross-flow system.40 The dimensions of droplets are controlled by the properties of the droplet and continuous phases such as viscosity and interfacial tension,65 the geometry of the microchannels and the relative flow rates of the droplet and continuous phases.40 Droplets have the diameters in the range from tens to hundreds of micrometers, which corresponds to volumes from 1 fL to 10 nL, respectively.66 Figure 1.3 shows an exemplary MF setup and an optical microscopy image of droplets laden with Jurkat cells, which were produced in a flow-focusing

MF device. The device consisted of three inlet channels: one inlet supplied a fluorinated oil (a continuous phase), while two other inlets supplied cell suspension and sterile cell-free media.

Variation in the ratio of the flow rates of the cell suspension and the media enabled control over the concentration of cells in the droplets.64

The frequency of MF droplet generation can be as high as several KHz, however typically, for cell encapsulation significantly lower frequencies have been used in order to avoid high shear stress imposed on cells during the encapsulation process. The number of droplets (and corresponding microgel particles) produced per unit time can be increased without significant

broadening of their size distribution by combining several droplet generators working in parallel.67 Furthermore, droplets can be generated “on demand” by implementing a computer- controlled actuation system, which opens or closes a series of valves in the MF device.68

Viscosity of the droplet phase is an important experimental variable in the MF formation of microgels: the utilization of highly viscous liquids counteracts the formation of monodisperse droplets and under particular conditions, can suppress the emulsification.65 On the other hand, subsequent to emulsification, it is imperative to trigger rapid gelation of droplets in order to form microgels in situ (on-chip), thereby preserving the narrow distribution of sizes of the precursor droplets.

Despite its great potential, MF encapsulation of cells has a limitation: it is a stochastic process and the number of cells per droplet is generally fitted to a Poisson statistics as:19,64 14

k e P(k)  k! (1) where P(k) is the fraction of droplets expected to contain k cells, and λ is the average number of cells per droplet. A more detailed discussion of the importance of Poisson statistics for cell encapsulation in MFs is given in Chapter 4. Thus MF encapsulation offers control over an average number of cells per droplet (or per microgel) by varying the size of droplets, and/or the concentration of cells in the feed suspension. The variation in the number of encapsulated cells is particularly important in single-cell studies. For example, the emulsification of a cell suspension containing 2.5106 cells/mL yields 37% of droplets containing a single cell, if the droplet diameter is 100 µm, while the fractions of empty droplets and droplets containing more than one cell are 26 and 37%, respectively.64

Recently, several groups attempted to overcome the limitation of MFs to control the number of cells per droplet. It was realized that the presence of a cell at the front of the elongated jet of the droplet phase leads to the break-up of the jet and produces a cell-laden droplet.69 This approach resulted in greater than 70% of droplets containing a single cell, however, is was limited to droplets with dimensions similar to a single cell, and lacked control over encapsulation of more than one cell per droplet. Figure 1.4 shows an alternative strategy for the encapsulation of a single cell per droplet utilizing the fluidic streamlines caused by the flow of cells in microchannels.70 By using an earlier finding on the highly ordered flow of evenly spaced particles,71 the frequency of droplet generation and their diameters were carefully tuned to produce a number of droplets containing exactly one cell.

Another challenge in MF cell encapsulation is the time-dependent change in the surface properties of the microchannel walls. It has been observed by our group, as well as by other research groups, that the surface properties of microchannels change in the course of cell encapsulation, presumably, due to the cellular release of surface-active species such as proteins, carbohydrates, or cell debris. The deposition of these molecules on microchannel walls often leads to their wetting with the droplet phase and results in poorly controlled droplet generation.

This problem can be overcome by the chemical modification of the microchannels prior to the encapsulation experiments. For example, hydrophobization of polydimethylsiloxane using silanization techniques can be used to prolong the lifetime of MF devices used for cell encapsulation.19,21

In principle, subsequently to MF emulsification, cells can be cultured in droplets that are suspended in a non-miscible liquid, however this method is not suitable for long-term cell culture, because of the inability to supply nutrients and remove waste products from the cell- laden droplets.64,72,73

1.4.4 Encapsulation of Cells in Microgels

In comparison with MF encapsulation of cells in droplets, encapsulation of cells in microgels offers the ability to transfer cell-laden microgels into an aqueous culture medium for prolonged cell culture. The composition and the structure of microgels can be conveniently varied to mimic natural microenvironments, e.g., ECM. The ability to change in a controllable, high-throughput manner the physical and chemical properties of the hydrogel, by changing the relative amounts of gel components, paves the way for studies of the role of the properties of 3D microenvironments on cell fate.

The general requirements to microgels as 3D microenvironments for cell encapsulation include their mechanical and chemical stability in aqueous media, such as a buffer or cell culture media, 17

biocompatibility, and sufficient permeability. The transformation of the cell-laden droplets into microgels can be achieved by the chemical or physical means.41 Chemically mediated gelation is carried out by photoinitiated or redox-initiated polymerization and/or crosslinking reactions of chemically reactive molecules. Physical gelation of biopolymers (more suitable for cell encapsulation) occurs by thermosetting or via ionic crosslinking. A common example of ionic chelation-induced physical gelation is the crosslinking of alginate with Ca2+ ions, which chelate? to the carboxylic acid groups of the guluronic units of the alginate molecules.74 A typical polymer undergoing temperature-induced gelation is agarose: it forms solutions at physiological temperatures (37oC), but upon cooling, the physical entanglement of the polymer strands yields aggregates which form a gel network structure.

1.4.4.1 Microgels Derived from Synthetic Polymers

The utilization of synthetic polymers for MF encapsulation of cells allows for a straightforward chemical modification of the cellular microenviroment. Typically, the first step involves the emulsification of cell suspension, which contains a photoinitiator, a crosslinker, and a monomer or a pre-polymer. This step is followed by a crosslinking/polymerization reaction to form a microgel. Figure 1.5 illustrates gelation of cell-laden droplets, which is triggered by UV- initiated photopolymerization and /or crosslinking. Since the irradiation can cause cell damage or death, it is important to conduct photoinitiated polymerization under mild conditions, e.g., by using a uniform short-time illumination of cell-laden droplets and minimizing the irradiation time.75

Owing to reduced cell viability upon UV-irradiation, redox polymerization is considered to be a promising alternative method.76 For example, Figure 1.6 illustrates yeast cells encapsulated in

microgels derived from the hyperbranched poly(glycerol decaacrylate) (hPG14.5Dea) and

77 poly(ethylene glycol diacrylate) (PEG4.6Dia). Polymerization of these prepolymers was carried out overnight at 30 oC and a low (2 g L-1) concentration of the initiator ammonium persulfate, and yielded cell viability of approximately 30%.

Figure 1.6 Viability of yeast cells encapsulated into the hyperbranched poly(glycerol decaacrylate) and poly(ethylene glycol diacrylate) microgels with a concentration of 450 and 100 g L-1, respectively. The samples were stained with Syto 9® (green dye, staining both living and dead cells) and propidium iodide (red dye, staining dead cells only) and then observed using bright field and confocal fluorescence microscopy. Yeast cell-laden microgels gelled overnight at 30 oC (a) and after treatment with 70% ethanol to kill all encapsulated cells prior to staining (b). Green staining corresponds to living cells; red or yellow staining marks dead cells. Adapted with permission from reference 77. Copyright © 2011 Elsevier.

1.4.4.1.1 Microgels Derived from Natural Polysaccharides

Natural polysaccharides such as alginate, agarose, pectin, and chitosan have been used in MF cell encapsulation, because of their biocompatibility and the mild conditions required to achieve gelation (thereby preserving cell viability).

1.4.4.1.2 Alginate Microgels

Alginate is a linear polymer composed of β-D-mannuronic acid and α-L-guluronic acid residues, which gels due to the coordination of divalent ions such as Ca2+, Ba2+ or St2+ cations to the carboxylic acid groups of α-L-guluronic acid units.78

Because of the rapid gelation causing a dramatic increase in viscosity, the MF emulsification of alginate solutions of moderately high concentrations is challenging.56 Typically, the emulsification and gelation steps are time-resolved by using the methods of internal gelation, external gelation or the fusion of droplets containing an alginate solution and a solution of

21,41,79,80 CaCl2. In the internal gelation method, droplets of alginate solution contained CaCO3 nanoparticles, and the continuous oil phase contained acetic acid, which diffused to the alginate droplets. Increase in the acidity of the droplets led to the dissolution of CaCO3 and the liberation of Ca2+-ions, which triggered alginate gelation.21 This method produced microgels with a well- defined morphology and a narrow size distribution. Under optimized conditions, the mammalian cells (Jurkat cells) encapsulated within the microgels, exhibited a viability of 74.3%.

Interestingly, cell viability increased at a higher concentration of CaCO3 nanoparticles, which

2- was attributed to the buffering effect of the released CO3 ions to counteract the excess acetic acid.

An external MF gelation was utilized in the formation of cell-laden alginate core-shell microcapsules.81 Two hydrodynamically focused co-axial streams, a cell suspension in cell media (휶-MEM) and an aqueous alginate solution, were broken into droplets in a flow-focusing

MF device. The droplets consisted of an outer alginate shell and a core comprising cell media and cells. The continuous phase contained a solution of CaCl2 in oleic acid. Diffusion of CaCl2 into the droplets led to the formation of a crosslinked alginate shell.

Figure 1.7 illustrates the encapsulation of cells in microgels formed by coalescence-induced gelation.41,82 Two distinct types of droplets were formed: one type of droplets contained human kidney cells in a 1.5 wt. % alginate solution in HEPES buffer saline, and the other type of droplets carried a 0.1M CaCl2 solution. The droplets coalesced in the downstream channel of the

MF device, thereby forming cell-laden alginate microgels. The viability of the encapsulated cells was 70%. This approach suffered from the lack of control over the coalescence of droplets, which resulted in polydisperse microgels and a limited control over the average number of cells/microgel.

A MF approach to the assembly of cell-laden alginate microgels can be further extended to the formation of microgels with a Janus structure.83-85 This method enables further manipulation cell-laden microgels. For example, a Janus particle composed of a calcium-alginate hydrogel was prepared with one half of the particle containing magnetic submicron particles, while the other half of the microgel contained encapsulated HeLa cells. The cell-laden particles were sensitive to an external magnetic field suggesting the possibility of using this method to spatially control the position of the microgel.

1.4.4.1.3 Agarose Microgels

Agarose is a linear polysaccharide consisting of alternating residues of β-1,3-linked-D-galactose and α-1,4-linked 3,6-anhydro-L-galactose. Aqueous agarose solutions form gels upon cooling, due to the aggregation of double helices formed by the physical entanglement of anhydro bridges on the individual molecules.86 Solutions of low-gelling temperature agarose are particularly useful for cell encapsulation: at moderately high concentrations: they are liquid at a physiological temperature of 37oC (which enables their emulsification), below 20oC they gel and upon heating to 37 oC they remain gel-like.

Recently, several research groups have utilized agarose as a biopolymer for MF cell encapsulation.19,42,87 Two recent reports from our group used a MF platform for rapid, throughput generation of agarose-based cellular microenvironments with varying properties.19,42

The suspension of cells in agarose solution was supplied to the MF droplet generator as two streams with various compositions. By varying the relative flow rates of the two streams but maintaining the total flow rate of the droplet phase constant, the composition of the precursor droplets was varied in a through-put manner, thereby enabling the generation of libraries of cell- laden agarose microgels with varying properties. Control over the mechanical properties of the agarose host was achieved by varying the concentration of the polymer within the microgels.42

The introduction into an MF device of two identical solutions of agarose, each containing different cell populations, was utilized to prepare cell-laden microgels with varying ratios of two cell types.19 Figure 1.8 illustrates a representative image of “red” and “green” murine embryonic 23

stem cells encapsulated in agarose microgels. Following the encapsulation of the cells in the microgels and their subsequent transfer to culture media, cell viability was 79.6 ± 2.5% and embryoid bodies were formed 4.5 days after the encapsulation.

1.4.4.2 Microgels Derived from Proteins

Cross-linked proteins have received much attention as potential materials for the generation of cellular microenvironments, because their chemical composition and fibrous structure mimic the properties of ECM, allowing cells to adhere, grow, move and differentiate.88

1.4.4.2.1 Gelatin Microgels

Gelatin consists of a mixture of amino acids, in which glycine, proline, and hydroxyproline are present in the most abundance. In aqueous solvents, gelatin gels via a cold-setting mechanism.89

This process is slow: at the temperature of ice water it may require up to several hours to form a moderately strong gelatin gel.90 Furthermore, in comparison with agarose, upon heating to physiological temperatures, a hydrogel derived from gelatin liquifies. The use of chemical crosslinking agents, e.g. glutaraldehyde may be problematic because of their cytotoxicity.91

The problem of liquification of cell-laden gelatin microgels produced by the MF method has been overcome by chemical modification of the gelatin prior to emulsification.20 First, unmodified gelatin microgels laden with rat adipose-derived stem cells were generated by emulsifying a suspension of the cells in an aqueous gelatin solution, followed by gelation of the droplets in an ice bath. The cell-laden microgels were then resuspended in an aqueous solution of phenolic hydroxyl-modified gelatin (gelatin-Ph). The cell-laden gelatin microgels were then encapsulated in droplets of gelatin-Ph which was subsequently gelled by both thermally-

mediated and peroxidise-catalysed crosslinking. The resulting core-shell microgels withstood heating to 37 oC. A layer of mouse L929 fibroblast cells was seeded on the surface of the cell- laden core-shell microgels. Figure 1.9a-c shows optical images of the core-shell microgels loaded with rat adipose-derived stem cells carrying a layer of fibroblast cells on their surface.

Figure 1.9d shows L929 cells seeded on the surface of the cell laden gelatin microgels.

1.4.4.2.2 Puramatrix Microgels

Puramatrix (SAPs or RADA 16) is a mixture of sixteen peptides, dissolved in salt-free water at concentration of 1 w/v % and pH=3. When RADA 16 solution is mixed with a solution containing salt, such as physiological media, the peptides spontaneously form a hydrogel.

The use of RADA 16 was demonstrated as a pre-gel agent for MF emulsification.92 Prior to emulsification, the cross-linking agent, finely powdered Dulbecco's Modified Eagle Medium

(DMEM) was dispersed in the continuous phase (Mineral Oil). Following the formation of cell- laden RADA 16 droplets, the powdered DMEM diffused into the droplets where it dissolved and crosslinked the droplets to form a cell-laden microgel using an external gelation approach. This method provided a method to encapsulate cells with minimal loss in cell viability.

Approximately 93% of bovine carotid artery endothelial cells showed survival, migration and growth in the microgels.92

1.4.5 Applications of Cell-Laden Microgels

Currently, applications of MF methods have mostly been reported for cell-laden droplets.

Microfluidic encapsulation of cells in droplets was used to create cell arrays for single-cell bioassays, including measurements of single-cell respiration rates,93 drug screening on the scale of a single cells,94 viability studies with different microenvironments,95,96 monitoring of cellular gene expression,97 and intercellular interactions,98,99 that is, single cell vs. cell-cell or cell subpopulations.

Encapsulation of cells in microgels offers the ability to transfer cell-laden microgels into an aqueous culture medium for prolonged cell culture. In addition, the composition and structure of microgels can be conveniently varied to imitate the natural microenvironments and in this manner, to study the role of properties of 3D microenvironments on cell fate. Microfluidic encapsulation of cells in microgels offers a unique ability to generate vast libraries of cellular microenvironments with a broad range of compositions and physical properties. The approach is illustrated schematically in Figure 1.10 for the microgels derived from agarose. In the first

strategy developed for co-encapsulation of different cell lineages (Figure 1.10a), two cell suspensions in agarose solution are supplied into the MF droplet generator.19 For the similar cell concentrations in two streams and the constant volumetric flow rate of the droplet phase, the ratio of the volumes of the two suspensions in the droplet is determined by the ratio of the respective flow rates. Thus, the variation in the flow rate ratios of the two cell suspensions enables control over the ratio of the co-encapsulated cells. This approach allows for the generation of a large number of microenvironments for cell co-culture, which can used for interrogation of cellular interactions.

The second strategy targets the generation of cellular microenvironments with varying mechanical properties.42 In Figure 1.10b, two cell suspensions, each in the solution with a different concentration of agarose (Cag, 1 and Cag, 2) were supplied to the MF device. The total flow rate of the droplet phase was maintained constant, whereas the ratio of the respective flow rates, Q1 and Q2, of the two suspensions changed. Following mixing and emulsification of the mixed cell suspension, the resulting cell-laden droplets were gelled by cooling. The microgels contained a varying amount of agarose, and hence, exhibited varying elasticity. A similar strategy can be used to produce cellular microenvironments with varying chemical compositions

(Figure 1.10c). Small molecules, DNA and various drugs can be introduced in different amounts in cell-laden agarose microgels.

The strategies shown in Figure 1.10 can be extended to the utilization of more than two streams of cell suspensions. Following cell culture in the microgel interior, cell fate can be analyzed by optical and fluorescence microscopy. The latter method, generally, requires the cells to be fluorescently labelled prior to encapsulation. An alternative approach is to lyse the hydrogel

matrix to release the cells and allow for detailed cellular studies using high through-put methods such as flow cytometry.19

A MF formation of cell-laden droplets (and consequently, microgels), followed by their transfer and cell culture in a second MF device provides the ability to study the dividision and growth of individual cells in a constant environment with the ability to control ambient conditions (pH, salinity, nitrogen concentration).72

Figure 1.10 Microfluidic generation of microgel-based cellular microenvironments with varying internal compositions. (a) Co-emulsification of two cell suspensions to achieve control over the ratio of co-encapsulated `red` and `green` cells by varying the volumetric flow rates of the suspensions, QR and QG, respectively; (b) Control of the mechanical properties of cell-laden microgels, achieved by varying agarose concentration in the microgels. Two cell suspensions in solutions with different concentration of agarose, Cag,1 and Cag,2 were supplied to the MF device and the ratio of the respective flow rates, Q1 and Q2, was changed in a throughput manner. (c) Two agarose solutions, one containing a suspension of cells, and the other containing a bioactive reagent are supplied in the MF device at varying flow rates Q1 and Q2. Copyright © 2012 WILEY-VCH.

1.4.6 Future work in Microfluidic Encapsulation of Cells in Microgels

Although encapsulation of cells in microgels is a very promising area of research in polymer materials science, MFs, and cell biology, currently, it faces several challenges. First, there is a

limited ability to controllably encapsulate one cell per droplet (and the resulting microgel). New, creative solutions are needed to exceed Poisson’s distribution in cell encapsulation, without compromising the distribution in microgel dimensions and the reproducibility and robustness of the MF method.

Second, the formation of libraries of cellular microenvironments often relies on the simultaneous change in several properties of microgels. For example, increase in microgel stiffness by increasing polymer concentration will concurrently change microgel pore size, fiber structure, and local polymer deformability, which will influence cell motility. Decoupling of the different properties is a significant challenge, which should be addressed through collaborations of cell biologists and polymer scientists. Furthermore, the encapsulation of cell suspensions critically depends on the viscosity of droplet phase and its interfacial tension, which can change in the presence of cells, addition of additives, or increase in polymer concentration. For example, increase in polymer concentration above a threshold value leads to a strong increase in solution viscosity and suppresses MF emulsification. Thus other methods of the variation in the mechanical properties of the microgels have to be developed.

Third, mixtures of biopolymers have been widely used as 2D substrates for cell culturing and scaffolds for tissue engineering.100 The use of multi-component systems is enticing: properties of one component can compensate for disadvantages of another, while the advantages of each individual component can be simultaneously realized. The properties of multicomponent gels, such as chemical composition, porosity, stiffness, elasticity, structural integrity, and cell adhesion can be tuned by varying the concentrations of appropriately chosen biopolymers.

Encapsulation of cells in microgels of biopolymer mixtures of collagen-agarose,57,101 and

chitosan-fibrin102 have carried out using homogenization technique. While there are many examples of the utilization of composite microgels for cell encapsulation, very few of them use a

MF platform.103,104 To our knowledge, there have been no reports of on-chip mixing of biopolymer solutions in order to tune the composition of cell-laden microgels in a high throughout manner. Utilization of multiple inlets would allow for the simultaneous addition of multiple components such as cross-linkers, biopolymers, and biologically active molecules in a high- throughput manner, which would enable precise control over the composition of the generated microgels.

Fourth, MF experiments on cell encapsulation will become more robust, when the problem of microchannel wetting is addressed. New, efficient, yet, not cytotoxic surfactants are required.

Fifth, currently, analysis of encapsulated cells is primarily performed using optical methods, such as optical microscopy but it will become more efficient if expanded to high through-put techniques such as flow cytometry, through the degradation of the microgel beads and subsequent release of the cells. Alternatively, the generation of smaller microgels with dimensions below 40 µm would enable direct injection of the cell-laden microgels into standard laboratory flow cytometers and allow for the analysis of the encapsulated cells without the need to remove them from the microgels. Finally, innovation and improvements in MF platforms will allow for the encapsulation, cell culture, and cell characterization using integrated probes, all conducted on a miniaturized lab-on-chip.

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Chapter 2 Materials and Methods

This chapter describes the materials and methods used in the present research from Chapter 3-4, and 6-7.

2.1 Materials

2.1.1 Biopolymers

Ultralow gelling agarose was purchased from Lonza (120 kDa, gel temperature 8-17oC,

SeaPrep, Swizterland). Phosphate Buffer Saline (PBS) and Hank’s Buffer Saline (HBSS) buffers were acquired from Gibco-BRL (Gibco-BRL, Rockville, U.S.A.). Gelatin was derived from bovine skin and was purchased from Sigma-Aldrich (Canada). Mineral Oil (30 cPs) and the oil soluble surfactant Sorbitan monooleate 80 (Span 80) were purchased from Sigma-Aldrich

(Canada). All biopolymer solutions were filtered and sterilized using syringe filters prior to use

(Millex GP, 0.22 µm pore size, Millipore, Canada)

2.1.2 Gases

Gaseous N2 (purity 99.99 %) and CO2 (purity 99.8 %) were purchased from BOC Canada.

Deionized water was purified by using a Mlli-Q Plus purification system (Millipore Corp.). NaCl was purchased from Sigma-Aldrich Canada. Dimethyl ether of poly(ethylene glycol) was donated by Dow Chemicals (Canada). Dr. Bridget Bergquist (Department of Geology, University of Toronto) kindly provided ocean water extracted from Bermuda coastal waters. Triton X-100

(TX-100) was purchased from Sigma-Aldrich, Canada and used as received.

2.1.3 Low Molecular Weight Compounds

Tris(hydroxymethyl)aminomethane (Tris) was supplied by Sigma Adlrich (Canada). Buffers were prepared by dissolving Tris in deionized water to achieve pH = 11.2. A 50 mM Tris buffers with pH < 11.2 were prepared by titrating them with an aqueous solution of HCl (pH = 2, EMD

Canada) at room temperature. Sodium hydroxide (NaOH) was purchased from EMD Canada.

Methyl methacrylate (MMA, 99%) was purchased from Fluka Canada and used without further purification.

2.1.4 Synthetic Particles

The synthesis of 2.8 μm-diameter poly(styrene-co-acrylic acid) (PS-co-PAA) particles was carried out as described elsewhere.1,2

2.1.5 Microfabrication

Poly(dimethylsiloxane) (PDMS, Sylgard® 184) and photoresist resin (SU-8 50, 3050) were purchased from Dow Corning (USA), and Microchem Co. (MA, USA), respectively. Silicon wafers were purchased from Wafer World Inc. Polycarbonate (CO) and cyclic co-olefin (COC) were purchased from Sabic Polymershapes and Zeonor Chemicals, respectively.

2.2 Methods

2.2.1 Mask Design

Masks were designed using AutoCad 2007® (Autodesk Inc., USA) software. The designs were printed on transparencies (Pacific Arts and Designs Inc., Markham, Canada) using ultra-high resolution printers with a resolution of 20,000 dpi.

2.2.2 Microfabrication of Negative Masters

2.2.2.1 Silicon Masters

Masters were prepared in a level 6 cleanroom facility (Bahen Center) at the University of

Toronto. Fabrication of the negative masters with microchannel features was carried out with

SU-8 photoresist (SU-8 50) on 3-inch diameter silicon (Si) wafer using photolithography.3,4 For the fabrication of microfluidic devices, the substrates were washed several times with acetone and methanol. The surface of the substrate was dried by blowing dry N2 gas. For Si wafers, spin- coating with SU-8 photoresist at a typical spin rate of 1200 rpm enabled the formation of 120 µm height features. After the spin-coating process, the photoresist layer was thermally baked to evaporate the solvent, γ-butyrolactone. Silicon wafers required only a single spin-coated layer of the photoresist prior to the generation of the microchannel patterns. Copper substrates require an initial seed layer of photoresist, followed by a second layer of photoresist which will be used to pattern the microchannel design.

A mask was placed overtop the photoresist layer on the Si wafers, and the system was exposed to

UV-light (Karl Suss MA6 mask aligner, λ=365~405 nm) to cure the photoresist and create the patterned microchannels. A typical exposure time was approximately 50 sec. Following UV- exposure, a post-bake process was conducted to enhance the crosslinking of the UV-exposed areas of the photoresist. After the initial exposure of the seed layer on the copper substrate, the second spin-coated layer was exposed through a mask to UV light to form the microchannel patters. The post-baked wafers with the crosslinked patterns of the microchannels were immersed for ca. 10 min in a developer solution (1-methoxy-2-propanol acetate) to remove the non-crosslinked regions of the photoresist. The substrates with the microchannel patterns were rinsed several times with isopropanol and methanol and dried under a gentle stream of nitrogen.

2.2.2.2 Copper Masters

The fabrication of copper masters was similar to the fabrication of Si masters. For copper wafers, spin rates of 3000 rpm led to 50 µm height features. An initial layer of SU-8 3050 was spin- coated onto the Copper substrate at a spin rate of 3000 rpm which lead to a layer of SU-8 with a height of 50 µm. Subsequent to photo-polymerization of the initial layer of SU-8, a second layer of SU-8 3000 was spin coated on top of the first layer to achieve a second 50 µm layer. The second layer was irradiated with UV-light through a pre-designed mask. Development of the master was carried out in a similar protocol to silicon masters.

2.2.3 Fabrication of Microfluidic Devices 2.2.3.1 Fabrication of PDMS Microfluidic Devices

Microfluidic (MF) devices were fabricated in poly(dimethylsiloxane) (PDMS) elastomer using a standard soft-lithographic procedure.5 The PDMS elastomer was prepared from Sylgard® 184

(Dow Corning Cop., USA). The Sylgard® 184 base polymer contains vinyl-terminated dimethylsiloxane oligomers, a platinum catalyst, and a silica filler (dimethylvinylated and trimethylated silica).6 The base polymer was mixed with the curing agent containing a crosslinker (dimethyl-methylhydrogen siloxane) and an inhibitor (tetramethyltetravinyl cyclotetrasiloxane) at 10:1 (w/w) ratio. Air was removed from the mixture under vacuum for 20 min. The mixture of the prepolymer mixture was poured onto the master and baked at 75 °C in an oven for more than 12 hrs. After curing, the replica was peeled from the master, and holes were created with a blunt-tipped needle at the designated positions, which corresponded to the inlets and outlets of the MF device. The replica and a substrate (a plane PDMS sheet) were oxidized in a plasma cleaner chamber (PDC-3XG, Harrick, USA) for 90 sec. The plasma-treated replica and the substrate were brought in contact and sealed immediately.

Polytetrafluoroethylene tubing (0.02 inch outer diameter) (Small Parts, USA) were forced tightly into the holes.

2.2.3.2 Fabrication of Thermoplastic Microfluidic Devices

Thermoembossing was conducted in a hydraulic press (Model 3851-C Carver Inc., Wabash, IN) with +/- 1oC temperature control of the top and bottom platens. The master fabricated on a copper substrate was loaded with its features facing up onto the bottom platen of the press. A 1 mm-thick sheet of the thermoplastic polymer was placed on top of the imprint template. It should be noted that polycarbonate exhibits a relatively high water absorption and requires a dehydration step, which involves pre-heating of the plastic at ~ 80oC for 40-80 min before embossing. A square polished copper plate with dimensions 7.6 x 7.6 cm and the thickness of 1 mm was placed on top of the polymer sheet, with the polished side against it.7

2.2.4 Microfluidic Experiments 2.2.4.1 Microfluidic Generation of Cell-Laden Agarose Microgels

Two cell suspensions were prepared in 2 wt% solutions of ultra-low gelling temperature agarose in PBS buffer pre-heated to 37oC. The concentration of cells varied in the range from 2 x 106 to 8 x 106 cells/mL. The suspensions with equal concentrations of cells were supplied to two channels of the MF device using two independently controlled syringe pumps (Harvard Apparatus 33

Dual Syringe Pump, U.S.A.). The temperature of the cell suspensions was maintained at 37oC in a temperature controlled incubator in order to prevent agarose gelation. The suspensions were mixed in a serpentine channel with a length of 250 mm. Mineral oil containing 3 wt% of the non- ionic surfactant Span 80 was supplied to the main channel of the MF device. Control over the encapsulation ratio of different cell populations was achieved by tuning the flow rate ratio of the individual suspensions, while maintaining the total flow rate of the two cell suspensions 43

constant. Gelation of the agarose droplets was induced by cooling them in the outlet tubing which was enclosed in a jacketed hose connected to a water circulator, which was using a 1:4 vol mixture of glycerol and water cooled to 2oC. The droplets passed through the tubing within ~ 5 min and were subsequently collected in a 15 mL centrifuge tube containing HBSS. The centrifuge tube was kept in an ice bath to complete the gelation of the microgels. After the microgels were maintained for 45 min in the bath, the suspension was centrifuged, the cell-laden microgels were separated, washed twice with HBSS buffer and transferred into HBSS buffer containing 2% v/v FBS. Collected samples were centrifuged at 1000 rpm at 4oC for 5 min. The centrifuged samples were washed with HBSS buffer and transferred to cell culture media.

2.2.4.2 Cell Culture 2.2.4.2.1 Mouse Embryonic Stem Cells

Mouse embryonic stem (mES) cells were cultured under sterile conditions and maintained in a

o 5% CO2 humidified incubator at 37 C. R1 and YC5-YFP-NEO, mES cells were maintained in cell culture media composed of Dulbecco’s Modified Eagle Media (DMEM, Gibco-BRL) supplemented with 15% (v/v) Fetal bovine serum (FBS, Gibco-BRL), 2mM L-glutamine, 0.1 mM beta-mercaptoethanol (BME, Sigma, St. Louise, MO), 0.1 mM non-essential amino acids

(NEAA, Gibco-BRL), 1 mM sodium pyruvate (Gibco-BRL), 50 µg/mL penicillin (Gibco-BRL),

50 µg/mL streptomycin (Gibco-BRL), and 500 pM Leukemia inhibitory factor (LIF, Chemicon,

Temecula, CA). Confluent dishes of mES cells were fed and passaged on alternate days.

Agarose microgels laden with mES cells were transferred to culture media comprising DMEM supplemented with 15% (v/v), FBS, 2mM L-glutamine, 0.1 mM BME, 0.1 mM NEAA, 50

µg/mL penicillin, 50 µg/mL streptomycin. Samples were maintained under sterile conditions and

o maintained in a 5% CO2 humidified incubator at 37 C for 4.5 days. 44

2.2.4.2.2 Human Megakaryoblastic Leukemia Cells

M07e and MBA2 human megakaryoblastic leukemia cells were maintained in cell culture media composed of Iscoves Modified Dulbecco’s Media (IMDM, Gibco-BRL), 10% (v/v) FBS, 50

µg/mL penicillin, 50 µg/mL streptomycin, 2mM L-glutamine, and 0.1 mM BME. Cells were passaged every three days (1:6 split ratio) and M07e cells were supplemented with 0.25 ng/mL

IL-3. Samples were maintained under sterile conditions and maintained in a 5% CO2 humidified incubator at 37oC.

2.2.4.2.3 Human Umbilical Cord Blood Cells

Umbilical cord blood cells were obtained from consenting donors according to procedures accepted by the ethics board of Mt. Sinai hospital (Toronto, ON, Canada). Mononuclear cells were processed as reported elsewhere.8 Progenitor cells were isolated from the mononuclear cell fraction using the EasySep human hematopoietic progenitor cell enrichment kit (Stem Cell

Technologies, Vancouver, BC, Canada), following the manufacturer’s protocol. Lineage- depleted cells (lin-) were cultured in serum-free conditions, as previously described8 for six days to achieve required cell numbers. Samples were maintained under sterile conditions and

o maintained in a 5% CO2 humidified incubator at 37 C.

2.2.4.2.4 Acute Myeloid Leukemia Cells

OCI-AML2 (AML2) is a human leukemia cell line established from peripheral blood of a patient with acute myeloblastic leukemia. AML2 cells grow in suspension as single round cells. The cells were maintained in cell culture media composed of Dulbecco’s Modified Eagle Media

(DMEM, Gibco-BRL) supplemented with 15% (v/v) Fetal bovine serum (FBS, Gibco-BRL).

Cells were passaged every four days (1:6 split ratio). Samples were maintained under sterile

o conditions and maintained in a 5% CO2 humidified incubator at 37 C. 45

2.2.4.3 Microfluidic Generation of Monodisperse Bubbles in Temperature Controlled Zones

CO2 gas was supplied to the MF device under a pressure PCO2=7psi (48.3kPa) or PCO2=12psi

(82.7kPa) through a polytetrafluoroethylene tubing (Small Parts, USA), attached to a Bellofram pressure regulator. A continuous liquid phase at a temperature of 23 +/-1oC was introduced into the MF device at the flow rate Qc=2.5 mL/hr using a syringe pump (Harvard Apparatus, USA,

PHD 2000 series). Various areas of the MF device were cooled by placing underneath them a hollow copper block and purging through it a glycerol/water mixture (1:4 v/v) at a temperature of 0oC. The temperature of the mixture was controlled using a water circulator (Neslab RTE-

040). A region of the MF device was heated by placing beneath it a 1.5 cm2 heating module (TE

Technology). The temperature in the module was controlled using an electronic temperature controller (TE Technology). The temperature of the cooling block and the heating elements were set to 0 and 35oC and the system was equilibrated for 30 min. The temperatures of the liquid phase were measured by using a thermocouple (TFCY-003, TFAL-003, Omega Engineering, 160

µm diameter) inserted into the microchannel.

Simulations of the temperature of the liquid travelling through the MF device were performed in the commercial Software Comsol (Comsol 4.0a, Comsol Inc. USA).

To measure the value of pH of the aqueous liquid phase, an in-flow pH meter (Cole-Parmer, 50

µL internal volume electrode, Ecomet P25 pH meter) was connected to the serpentine channel at distances of 135, 270, and 390 mm from the orifice.

2.3 Characterization

2.3.1 Optical Microscopy Imaging

An Olympus BX41 or BX51 microscope (Olympus, USA) with a high-speed camera

(Photometrics CoolSNAP ES) was used to image the generation of bubbles and droplets in the

MF device. The same setup was also used to image bubbles and droplets on a glass slide. Optical microscopy imaging of the CO2 bubbles moving with the continuous phase was carried out in reflection mode. In order to acquire images, a silicon wafer was placed between the MF device and the cooling copper block or the heating module.

2.3.2 Size Distribution of Bubbles and Droplets

The dimensions of bubbles, droplets and particles generated by MF means were analyzed using

Image Pro Plus (Media Cybernetics, USA) software. The polydispersity of bubbles or droplets was characterized as the coefficient of variance CV (%) = (σ/D)x100 where σ is the standard deviation of the size of bubbles or droplets and D is the mean diameter of bubbles or droplets.

The volume, V, of spherical bubbles was calculated as V = (4/3)π(D/2)3. When the value of D exceeded the height, h, of the microchannels, the bubbles acquired a discoid shape and their volume was approximated as V = (π /12)[2D3-(D-h)2(2D + h)].9

2.3.3 Characterization of Cell-Laden Agarose Microgels

Fluorescence microscopy images of cells labeled with a fluorescent dye, were captured using a

Zeiss Microscope (Axio Observer D1, U.S.A.) coupled with a digital camera (Axio Cam HRm,

Zeiss, U.S.A.). Ratios between different dye-labeled encapsulated cells were determined by analyzing images obtained by optical fluorescence microscopy. Approximately 8000 cell-laden

microgels were included in the analysis, in order to determine the optimized conditions for cell encapsulation.

2.3.4 Flow Cytometry 2.3.4.1 Analysis of Cell-Laden Agarose Microgels

Encapsulation ratios of R1 mES cells were monitored using a Biosort flow cytometer (Union

Biometrica, U.S.A.) with a bore diameter of 250 µm. The cytometer was equipped with 488 and

561 nm excitation lasers. Prior to encapsulation, the cells were labeled with CFDA SE (green) or

CMTMR (red) dyes (Invitrogen). Microgel samples were introduced into the flow cytometer following the manufacturer’s instructions (Union Biometrica).

The sorting process was performed according to the manufacturer’s instructions (Union

Biometrica). The cell-laden microgels were gated to exclude 1.5% of cell-free microgels and cell debris by using the gating parameters Time of Flight vs. Extinction. Time of Flight provided the time required for an object to pass through the instrument detector, that is, the length of an object. Extinction reflected the internal complexity of an object similar to side scatter in conventional flow cytometry instruments. Microgels were then sorted based on fluorescence intensities, that is, Red Peak Height vs. Green Peak Height. We set the parameters for the fluorescence signal amplification as follows: Full scale: Tof 256, Ext 256, RedPH 65536,

GreenPH 65536; Gains (Signal): Ext 3, Green 2, Red 2; Trigger: Ext Thresholds: Signal 30, TOF

Minimum: 10; PMT Control: Green 500, Red 600.

2.3.4.2 Analysis of Single Cell Viability

M07e cells labeled with CFDA SE were monitored by fluorescence-activated cell sorting

(FACS). Following 4.5 days incubation of the cell-laden microgels in a co-culture environment,

agarose microgels were digested by adding a 0.1 % v/v aqueous solution of agarase enzyme

(Sigma) and maintaining the system for 30 min at 37oC. In the control experiments conducted with non-encapsulated M07e cells it was established that no significant change in cell viability occurred before and after the digestion of agarose with agarase (82.3±1.5% and 81.8±1.1%, respectively). The released cells were washed with HBSS buffer and centrifuged at 1000 rpm for

5 min at r. t. to remove excess agarose. 7-Aminoactinomycin D (7-AAD) dye (Molecular

Probes, U.S.A.) was added at 1 µg/ml to cell suspensions to label non-viable cells. The dispersion of cells was transferred to an appropriate tube for FACS analysis and the viability of

M07e cells was determined as a function of the encapsulation ratio of the MBA2 and M07e cell populations.

2.3.4.3 Analysis of % Rescue of UCB cells

Flow cytometry analysis was performed on cells prior to their encapsulation in microgels, using a FACSCanto flow cytometer (BD Biosciences, San Jose, CA, USA), to assess for cell type frequencies. Cells were stained with the following conjugated antibodies: CD34-APC (BD

Biosciences), CD133-PE (Miltenyi Biotec, Auburn, CA, USA), CD14-PE (BD Biosciences),

GyA-PE (BD Biosciences), CD41-PE (BD biosciences), and CD11b-APC (BD Biosciences). 7-

AAD dye was added to assess cell viability and isolate live cells for quantification. UCB cells were then encapsulated in microgels either in the absence of MBA2 cells (1:0 ratio), or in the presence of MBA2 cells in a 1:1 ratio. MBA2 cells were labeled with CFDA SE dye prior to encapsulation, in order to distinguish MBA2 cells from UCB cells in subsequent analysis. The cells encapsulated within the microgels were cultured in Iscove’s Modified Dulbecco’s Medium

(IMDM, Gibco) with no addition of growth factors or serum substitutes. Following three days of co-culture incubation, the agarose was digested, counts of viable cells were performed, and flow

cytometry analysis was repeated. Cells labeled positive in the FITC channel were excluded to remove all MBA2 cells from analysis, and viable hematopoietic cell sub-populations were quantified as:

#Viable Cells (3days post  encapsulation) %Re scue  x100% #Viable Cells ( pre  encapsulation) (1)

2.3.4.4 Analysis of the Variation in Cell Numbers of AML2 Cells

20 µL cell-laden macroscopic agarose gels were generated by preparing agarose solutions at concentrations of 1, 2, 3, 4, and 5 wt. % loaded with AML2 cells. The concentration of cells in all samples was 1.25x105 cells/mL. 20 µL aliquots of the agarose/cell suspension were transferred into individual wells of a 96-well plate. Triplicate samples of each concentration of agarose were prepared. Positive controls consisted of: (i) agarose gels without cells for each concentration of agarose, and (ii) AML2 cells suspended in media without the presence of agarose gel. The 96-well plate was stored at 4 oC for 45 minutes to induce gelation of the agarose. Proceeding gelation, 100 µL of cell media was added to each well and the samples were

o incubated for 7 days in 5% CO2 at 37 C. On day 7, 10 µL of CCK-8 dye (Dojindo Molecular

o Technologies Inc., Japan) was added to each sample and allowed to incubate in 5% CO2 at 37 C for 3 hours. The 96-well plate was then placed in a UV/Vis plate reader (SpectraMAX

GeminiXS, SOFTmax Pro). Absorbance measurements at 450 nm (휆max for CCK-8 is 450 nm) were acquired for all samples.

2.3.5 Characterization of Agarose Solutions 2.3.5.1 Characterization of Rheological Properties of Agarose Solutions

The viscosity of agarose solutions with various concentrations of agarose was determined using a rotational cylinder rheometer (Brookfield Model VD-III, Enhanced UL Adapter). Temperature was maintained at 37 oC using a temperature controlled water circulator (Fischer Scientific,

Isotemp 3016H). In viscosity measurements, agarose solutions were inserted into the temperature controlled sample holder at 37 oC. The viscosity measurements were conducted in the Newtonian regime by adjustment of the spindle spin rate until the viscosity of the agarose solution remained constant.

2.3.5.2 Measurement of the Elastic Modulus of Agarose Gels

The elastic modulus of agarose gels was measured using two independent techniques.

Macroscopic gels were measured using an Instron 5848 Microtester with a 50N load cell. The agarose gel was removed from a PBS buffer and cylinders with a diameter of 18 mm were cut from it. The sample was loaded onto the microtester and compressed at a rate one-tenth of its height in millimeters per minute, and held to allow for stress relaxation until equilibrium was reached. The Young’s moduli of the gels were calculated from the ratio of the recorded stress at equilibrium and the applied strain. The measurements were conducted at r.t.

In comparison, agarose microgels with different concentrations of agarose were measured by

Atomic Force Microscopy (AFM) at r.t and at 37 oC. AFM measurements on the agarose microgels were performed using modified gold coated silicon-nitride cantilevers. AFM measurements were conducted by placing a sample of agarose microgels dissolved in PBS buffer on a gold substrate. The tipless cantilever was lowered onto the microgel, and aligned with the center of it. A loading force was applied to compress a microgel at a rate of 0.2 Hz until a set 51

point with a certain magnitude of force was reached. For each microgel composition, a minimum of five microgels were examined by collecting 6 force-distance curves for each of gel. The force indentation results were fitted to the following modified Hertz law equation using Igor Pro software:10

(2)

α is the indentation distance, P is the force (pN), σ is the Poisson’s ratio (which was assumed to be 0.5), E is the Young’s modulus (Pa), and D is the microgel diameter.

2.3.6 On-Chip Characterization of Chemical Reactions 2.3.6.1 Measurements of pH

The customized pH probe (MI408C, Microelectrode Inc.) and flow-through reference probe

(ME16730, Microelectrode Inc.) were connected to a pH meter (VWR Symphony SB70P). The calibration of the probe was conducted using pH 4 and pH 7 buffer solutions (Sigma Aldrich,

USA), as suggested by the manufacturer. The accuracy of the measurements was +/- 0.01 of pH unit. The pH probe was integrated into the MF device via threaded interconnects.

2.3.6.2 Measurements of Temperature

K-type thermocouples were fabricated from 80 μm-diameter chromel and alumel wires (TFCY-

003, TFAL-003, Omega Engineering). A heating pad (KHR-2, 10 Watt, Omega Engineering) was connected to a Proportional, Integral, Differential (PID) temperature controller (CN79022-

C4, Omega Engineering). On-chip temperatures were controlled using commercially available software (CN7A201, Omega Engineering). The thermocouple probe was integrated into the MF device via threaded interconnects.

2.3.6.3 ATR-FTIR Experiments

A Veretx 70 spectrometer (Bruker Inc.) was equipped with a deuterated L-alanine doped triglycene sulphate (DLATGS/D301, Bruker Inc.) and single bounce diamond ATR accessory

(MIRacle, Pike Technologies). A purge gas generator (75-45, Parker Balston) was used to limit absorption by ambient CO2 (g) and H2O (g). Opus 6.5 software was used for computer-control of data acquisition and analysis.

References

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(6) A. Olah, H. Hillborg, G. J. Vancso, Appl. Surf. Sci. 2005, 239, 410.

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Chapter 3

High-Throughput Combinatorial Cell Co-Culture Using Microfluidics

3.1 Introduction

Autocrine and paracrine signalling is paramount for both local and systemic cell-cell interaction networks, which modulate development and homeostasis and underlie both ageing and disease progression.1 In vivo, stem cell (SC) fate may be regulated by tissue- specific niches, such as the bone marrow, vascular, and hepatic microenvironments.2 The fate of stem cells is regulated in the niche through chemical and biological effectors, such as the release of cytokines, cellular contacts via adhesion molecules, and by interactions with components of the extra-cellular matrix.3 One of the key functions of SC niches is to maintain tissue homeostasis through direct and indirect cell-cell interactions. Homesostasis of SCs is achieved by balancing the fractions of quiescent and activated cells, and the signals they produce. Stem cell fate can be directed through the activation of specific cues within the niche to proliferate, self-renew, or differentiate to more specific cell types.4 In addition to gaining fundamental understanding of cellular interactions, directing cell fate by co-culturing specific cell lineages with SCs has potential applications in wound healing,5 tissue engineering,6 and cancer research.7 Cell co-culture is also important in establishing and maintaining embryonic SC cultures from both mouse8 or human sources, as well guiding the differentiation of SCs towards specific lineages. As an example, co-culture of OP9 stromal cells (derived from Macrophage-colony stimulating factor deficient mice) with embryonic

SCs leads to primitive and definitive hematopoetic development of the SCs in vitro, in a manner similar to that occurring in murine ontogenesis.9

The development of new and efficient methods which would provide the high-throughput generation of massive libraries of co-cultured cells has fundamental and practical importance. These methods should enable (i) the controlled generation of interactive cellular microenvironments that closely mimic the native cellular microenvironment in chemical and mechanical properties; (ii) the spatio-temporal control over the relative number of co- cultured cells in the microenvironment; and (iii) the ability to carry out high-throughput screening of the behaviour of co-encapsulated cells. Futhermore, since cell-cell interactions can occur through direct contact or through soluble factors via paracrine signalling, it is important to maximize the interaction between neighbouring cell types by creating microenvironments with an inter-cellular distance on the scale of cells.

Currently, micropatterning of planar substrates does not provide the ability to co-culture cells in 3D environments.10 Other approaches have been shown to be capable of producing 3D microenvironments for co-cultured cells, such as microfabrication of scaffolds11 or the generation of sphere-shaped cell colonies.12 Another approach includes the mixing of cell- laden and cell-free microgel modules at varying ratios to produce microscale constructs with a controllable ratio of the two types of modules, and hence the controllable concentration of cells in the overall construct.13 Yet, these approaches have a limited ability to vary the relative ratios of the co-cultured cells in a continuous high-throughput manner, and therefore are limited in their ability to create large libraries for rapid screening of paracrine signalling mediated cell-cell interactions.

Bulk emulsification of single cells or cell aggregates14,15 and more recently, microfluidic

(MF) emulsification of suspensions of cells has offered a new operational platform in cell biology.16-18 Rapid MF generation of thousands of cell-laden droplets with precisely

controlled dimensions and compositions provided the ability to create a large number of well-defined 3D microenvironments for studies of cell growth and viability, 19,20 gene expression,21 and enzymatic activity.22-24 Encapsulation of cells in hydrogel microbeads

(microgels) derived from precursor droplets, yielded biocompatible microenvironments, which allowed the exchange of nutrients and oxygen between the encapsulated cells and the surrounding medium.25-28 Microfluidic encapsulation of cells in droplets and microgels has paved the way for the high-throughput generation of combinatorial libraries of assays,29-31 however, microenvironments for co-cultured cells with a controlled ratio of the number of cells of different lineages has not been demonstrated until this work.

In the present chapter, a MF strategy is described for rapid (40 s-1) generation of 100 µm diameter (3D) agarose microenvironments for cell co-culture. Agarose is a neutral polysaccharide that undergoes gelation upon cooling.32 Agarose is bio-inert, it exhibits low adsorptivity to proteins and cells, it can be readily functionalized with cell adhesion proteins, and its mechanical properties can be tuned by varying the agarose concentration in the gel.33-36 We demonstrate the practical advantage of the MF method for the generation of cellular microenvironments by (i) achieving a 100% encapsulation efficiency; (ii) by growing pluripotent cell-derived aggregates in the microgels, (iii) by controlling the extent of paracrine signalling between M07e human megakaryoblastic leukemia cells and MBA2 support cells, which were co- encapsulated in varying ratios, and (iv) by co-encapsulating MBA2 cells in combination with heterogeneous umbilical cord blood cells to discover and determine the rescue effect of localized the localized growth factor (IL-3) on the various hematopoietic sub-populations. The impact of the ratio of the encapsulated cells, MBA2 to M07e and MBA2 to UCB case studies demonstrates a novel approach to the controlled generation and characterization of paracrine signalling

amongst cell-populations in an in vitro setting. In comparison with existing methods used for cell encapsulation, this approach allows for the continuous high-throughput generation of microenvironments with a tuneable ratio of encapsulated species, and the generation of large combinatorial co-culture libraries (>100,000 samples/hr).

3.2 Results and Discussion

Figure 3.1 a and b show an optical microscopy image and schematic illustration of the MF method for producing 3D hydrogel microenvironments for cell co-culture. Aqueous solutions of agarose carrying two distinct populations of cells were supplied at 37oC to the MF droplet generator (streams R and G) at a constant flow rate QTot. Prior to reaching the T-junction, mixing of streams R and G was enhanced by passing them through a serpentine channel

(Figure 3.1a). A stream of mineral oil (viscosity 30 cP) was supplied to the MF device at a right angle to the stream of the mixed cell suspensions. At the T-junction, the shear stress imposed by the stream of the mineral oil onto the stream of the mixed cell suspension led to the breakup of the agarose solution and the generation of droplets of the agarose cell suspension. During the encapsulation process, cells remained in the aqeuous suspension and were not released into the continuous phase.

The diameters of the droplets were tuned from 70 to 110 µm by varying the flow rate of the continuous oil phase, Qc, from to 1.2 to 2.5 mL/h (at QTot = 0.09 mL/hr). The relative standard deviation in droplet dimensions did not exceed 2.5 %. At the end of the downstream channel the agarose droplets and the mineral oil was cooled to 2 oC by encasing the outlet tubing in a temperature controlled water circulator tubing. Gelation of the agarose solution led to the transformation of droplets into agarose particles laden with cells (microgels). The microgels were collected in 15 mL centrifuge tubes containing HBSS buffer cooled to 4 oC. Uniform and

fast gelation of the droplets yielded microgels with a polydispersity below 3% (Figure 3.1c). The final size of the microgels in the HBSS buffer was approximately 10 % smaller than the diameter of the precursor droplets, owing to droplet shrinkage upon gelation. Cell-laden microgels were centrifuged, washed with the HBSS buffer and transferred to the appropriate cell media culture.

3.3 Controlling Encapsulation Efficiency

To achieve a 100 % encapsulation efficiency of cells within precursor droplets, such that each droplet contains at least one cell, we relied on the Poisson distribution for the compartmentalization of cells as in:37

e x P(x)  (2) x!

where P(χ) is the fraction of droplets expected to contain χ cells, and λ is the average number of cells per droplet. Since χ represents a specific number of cells per droplet, it must be an integer value starting from 0 (for droplets containing no cells).

We used a built in Poission Distribution function in Microsoft Excel (Microsoft Office 2007) to calculate the probability (a fraction) of droplets containing χ number of cells (P(χ)). The

Poission Distribution function in Microsoft Excel required the user to input a range of integer values for χ (we chose to use x in the range from 0 to 6. For each of the set values of

χ we calculated λ (calculated based on the concentration of cells in the agarose solution and the droplet diameter for a specific experiment) which was determined as the ratio between the concentration of cells in the droplet phase and the average volume of a droplet. Inputing the range of values of χ for a specific λ, the Poission Distribution function in Microsoft Excel generated a series of probability (or fraction) values, (P(χ)), which were plotted as a function of χ to generate the graphs shown in Figure 3.2.

Figure 3.2a displays the variation of the percent probability (fraction) of droplets containing a particular number of cells, χ, at varying 휆 (which was varied by changing the diameter of the droplets). Similarily, Figure 3.2b shows the variation in the fraction of droplets

containing a particular number of cells χ by varying the value of 휆 as a function of the initial cell concentration in the agarose solution.

By setting the concentration of cells to e.g., 8x106 cells/mL we used eq. (2) to establish that for 110 µm-diameter droplets, the encapsulation efficiency was 99.7 %, which was sufficiently close to a 100 % encapsulation rate.

In the encapsulation experiments, the concentration of cells in the agarose solution varied from 2x106 to 8x106 cells/mL. For a particular cell concentration, the encapsulation efficiency was controlled by tuning the diameters of droplets from 70 to 110 µm. Figure 3.3 shows that the theoretical and experimental results for the total number of cells per droplet

(without differentiating between the different types of cells) were in excellent agreement. For

example, for 70 µm-diameter precursor droplets and a cell concentration of 8x106 cells/mL, we achieved a 78.0 % encapsulation efficiency vs. the predicted value of 76.2 %, demonstrating concordance between theory and experiment.

An encapsulation efficiency of 98.5 % was achieved for 110 µm-diameter droplets at a cell concentration of 8x106 cells/mL. At a cell concentration of 2x106 cells/mL, a maximum encapsulation efficiency of 74.6 %, in agreement with a theoretical encapsulation efficiency of

75.2 %. Figure 3.4 shows an excellent correlation between experimental results of the number of cells per microgel and the theoretical values of the number of cells expected per microgel as determined by the Poisson distribution for each discrete number of cells per droplet. Experiments were conducted at cell concentrations of 4x106 and 8x106 cells/mL. The values of P(χ), χ, and λ 62

for these experiments are shown in Tables 3.1 and 3.2. Co-encapsulation of cells was tested for mES (murine embryonic stem) cells labelled with Vybrant CellTracer (CFDA SE “green”) and

CellTracker Orange (CMTMR “red”). Two suspensions, each containing red or green mES cells

(streams R and G, respectively), at a concentration of cells of 8x106 cells/mL, were supplied to the MF device at equal volumetric flow rates QR=QG=0.045 mL/hr. Figure 3.5a shows a representative image of red and green mES cells encapsulated in agarose microgels. Following the encapsulation of the cells in the microgels and their subsequent transfer of the microgels into culture media, cell viability was measured to be 79.6 ± 2.5 % and embryoid bodies were observed 4.5 days after encapsulation (Figure 3.5b). It should be noted that the viability of individual mESC cells in cell culture is typically in the range from 20 to 50 % and is dependent on the cell line used.38

In co-encapsulation experiments, the concentration of cells in each stream was 8x106 cells/mL and the total flow rate of the R and G streams was 0.09 mL/hr. The ratios of flow rates of the R- to-G streams were 1:0, 4:1, 1:1, 1:4, 0:1, where the flow rate ratios of 1:0 and 0:1 corresponded to the generation of droplets from a single stream (R or G, respectively), thereby encapsulating a single cell population.

We note that when the two cell suspensions were mixed, the suspensions of the individual cells were diluted in proportion to the relative flow rates. The effect of dilution on the encapsulation concentration of each cell population was described by the Poisson distribution expected for 110

µm diameter droplets and the corresponding cell concentration in the mixed suspension (Figure

3.6).

χ 0 1 2 3 4 5 6 λ D (µm) P(χ) 110 24.81 34.58 24.10 11.20 3.90 1.09 0.25 1.39 100 35.09 36.75 19.24 6.72 1.76 0.37 0.06 1.05 90 46.61 35.58 13.58 3.46 0.66 0.10 0.01 0.76 80 58.50 31.36 8.41 1.50 0.20 0.02 0.00 0.54 70 69.82 25.08 4.50 0.54 0.05 0.00 0.00 0.36

χ 0 1 2 3 4 5 6 λ D (µm) P(χ) 110 0.38 2.11 5.89 10.95 15.26 17.02 15.81 5.58 100 1.52 6.35 13.30 18.58 19.45 16.30 11.38 4.19 90 4.72 14.41 22.00 22.39 17.10 10.44 5.31 3.05 80 11.71 25.12 26.93 19.25 10.32 4.43 1.58 2.14 70 23.77 34.15 24.53 11.75 4.22 1.21 0.29 1.44

The encapsulation of red and green mES cells in the microgels was analyzed using optical fluorescence microscopy (Zeiss Axio Observer D1) and flow cytometry (Biosort, Union

Biometrica, 250 µm-diameter bore size) (Figure 3.7a-e). The utilization of flow cytometry provided a high-throughput approach to analyzing cell-laden microgels. In Figure 3.7, the encapsulation of red and green mES cells at various ratios is reflected by flow cytometry

graphs with populations in three separate gating regions, which were determined by the ratio of the red-to-green cells from the entire microgel population, that is, from the all the encapsulated cells, not from individual microgels. As expected, the samples comprising

microgels laden solely with green (R:G = 0:1) or solely with red mES cells (R:G = 1:0)

(Figure 3.7a and Figure 3.7e, respectively) were characterized by signals appearing exclusively in their respective gating regions. Signals appearing in the third region (double positive for both red and green cells) were a direct result of a red cell and a green cell residing in the same path length of the flow cytometer excitation laser. This effect results in the instrument detecting a double signal comprised of both a red and green component.

Signals in the double positive region could not be quantitatively assessed, due to the uncertainty in the number of red or green cells which contributed to the overlapped signal.

To quantitatively compare the results of flow cytometry experiments, only signals appearing exclusively in the red region or exclusively in the green region were compared (Flowjo 7.5).

Signals collected exclusively in the red-gated and the green-gated regions were used for analysis, and signals outside of either the red or green gate were excluded from the analysis.

The microgels generated at the 1:4 flow rate ratio of the R-to-G streams (Figure 3.7b) contained ~80 % of the cell population in the green gating region and ~20 % in the red- gating region. The microgels produced at equal flow rates (1:1 ratio) of the R and G streams

(Figure 3.7c) had statistically equal ratios of red-to-green cells of 47.7 % and 52.3 %, respectively, appearing in the pure red and pure green regions. The microgels formed at the

R-to-G flow rate ratios of 4:1 (Figure 3.7d) showed an encapsulation trend that was inverse to that shown in Figure 3.7b, with populations of red and green cells of 79.7 % and 20.3 %, respectively. The results of the analysis of cell co-encapsulation by flow cytometry and optical microscopy images were in excellent agreement (Figure 3.7f).

Figure 3.7 Co-encapsulation of mES cells in microgels achieved at varying flow rate ratios of the corresponding cell suspensions. (a-e) Left: Relative fluorescence intensity plots of red and green channels for sorted microgels laden with R1 mES cells labelled with Vybrant CFDA or CellTracker™ Orange CMTMR (green and red cells, respectively). Right: Optical microscopy images of the corresponding microgels. The microgels were produced at the respective flow rate ratios of the R and G streams: (a) QR:QG = 0:1, (b) QR:QG = 1:4, (c) QR:QG = 1:1. (d) QR:QG = 4:1, (e) QR:QG = 1:0. Gating was determined by positive controls comprising cells labelled with only one dye (R:G = 1:0 and 0:1). (F) Fraction, α, of green and red cells encapsulated at different ratios of QR:QG. Light and dark green bars show the fraction of encapsulated green cells, determined by image analysis and flow cytometry, respectively. Light and dark red bars represent the fraction of encapsulated red cells, determined by image analysis and flow cytometry, respectively. Fluorescence intensity scale is defined by the sorting parameters. Scale bar is 100 µm.

3.4 Co-encapsulation of MBA2 Cells and M07e Cells

A specific application of the MF cell co-culture platform was demonstrated by uncovering cellular interaction networks. We encapsulated in the microgels, at varying ratios the MBA2 cell line (secreting IL-3) and the IL-3-dependent M07e human megakaryoblastic leukemia

cell line.37 The M07e cells are a sub-line of the M07 human megakaryoblastic leukemia cell lines that require interleukin-3 (IL-3) for proliferation and survival. MBA2, a cell line transfected with the P210 gene, secretes IL-3 and is used in both an autocrine and paracrine fashion. We hypothesized that the survival and proliferation of encapsulated M07e cells could be controlled by varying the relative number of MBA2 cells in the microgels (Figure

3.8a).

M07e and MBA2 cells were co-encapsulated in the microgels at the following flow rate ratios (M07e:MBA2) = 0:1, 1:9, 1:4, 1:1, and 4:1. The viability of the M07e cells was determined after 4.5 days (Figure 3.8b). As expected, low survival of M07e was observed in the presence of a small number of MBA2 cells due to the lack of IL-3 present in solution.

With increasing fractions of MBA2 cells in the microgels, M07e viability increased, reaching a maximum of 65.9 ± 8.4 % at the ratio of the encapsulated cell lines of 1:1. To verify that higher M07e viability results from the higher relative number of MBA2 cells (therefore increasing the concentration of endogenous IL-3), we conducted control experiments with non-encapsulated M07e cells in the presence of exogenous IL-3. At an IL-3 concentration of

10 ng/mL, the viability of M07e cells was 76.3 ± 3.5 % after 4.5 days. This modestly higher value than that achieved for a 1:1 ratio of encapsulated MBA2:M07e cells was presumably caused by the higher levels of IL-3 in the control system.

Since M07e cells are a parental line to the MBA2 cells, standard approaches to distinguish the two populations by side scatter (SSC) vs. front scatter (FSC) could not be applied. The

SSC vs. FSC was utilized in distinguishing between viable and non-viable M07e cells, based on the change in cell morphology upon cell death. In order to differentiate between the two cell lines, M07e cells were marked with the CFDA dye prior to encapsulation in the

microgels. The dye was retained by the cells throughout cellular development, inherited by daughter cells, and was not transferred to adjacent cells.39-41 Furthermore, to quantify the viability of M07e cells after their encapsulation in microgels, 7-Aminoactinomycin D (7-

AAD) dye was added to all cell samples prior to FACS analysis. 7-AAD cannot readily pass through intact cell membranes and is commonly used for DNA staining of dead cells. The fluorescence intensity as a result of cellular uptake of 7-AAD was determined by flow cytometry.

Typical histograms of the cellular uptake of 7-AAD dye by CFDA-labelled cells at various co-culture ratios of M07e-to-MBA2 cells are shown in Figure 3.8c. Live cells were categorized as 7-AAD negative and dead cells were categorized as 7-AAD postive. A decrease in the M07e:MBA2 ratio (increase in the relative number of encapsulated MBA2 cells) resulted in a greater number of M07e cells appearing as 7-AAD negative in the flow cytometry histograms.

To verify that endogenously produced IL-3 resulted from cell co-encapsulation within the microgels, rather than diffusion of IL-3 through the medium, we carried out control experiments. We encapsulated either MBA2 or M07e cells in agarose microbeads and mixed these two cell-laden microgel populations in a 1:1 ratio in the varying volume of media.

Encapsulation of each cell line in the microgels was carried out at a cell concentration of

8x106 cells/mL. Figure 3.9 shows that regardless of the volume of the media (corresponding to the varying distance between the microgels), M07e cells had viability not exceeding 20 %.

We attribute this effect to the limited diffusion of IL-3 from the MBA2-laden microgels into the M07e-laden microgels, thereby resulting in low concentrations of endogenously produced

IL-3 in the bulk media.

3.5 Co-encapsulation of Heterogeneous Population of Primary Human Hematopoietic Cells with MBA2 Cells

An alternative application of the MF platform, we investigated the impact of a growth factor on specific cell types within the primary human hematopoietic SC culture system (Figure

3.10a). Cultures of hematopoietic cells are highly complex and heterogeneous. The heterogeneity is a result of the culture dynamics and paracrine signalling cascades that act to regulate the system. A study of the role of specific ligand-cell interactions within the context of this system would enable a greater understanding of signalling networks and provide a means of manipulating the culture system, however, currently this task is biologically and technically challenging. Figure 3.9 shows that cells encapsulated in each microgel are isolated from each other, therefore enabling high throughput (parallel) interrogation of interactions between two types of cells in the same microgel, but are unaffected from other

encapsulated cells in other microgels. An important aspect of this approach is the co- localization of two types of cells within a small sample volume. In comparison to bulk encapsulation in a macroscale agarose gel, the probability of having two types of cell populations in close spatial proximity to each other would be very low and highly variable, making it difficult to examine cellular interactions and their effects across a highly heterogeneous cell population in a controlled manner.

In our work, after co-culturing the IL-3-secreting MBA2 cells with heterogeneous UCB cells, the UCB cells were assessed for the specific effect of localized IL-3 secretion on the viability of a particular type of cell. By assessing the cell phenotype of the viable UCB cells, the rescue (relative to control conditions in which MBA2 cells were not present) of specific hematopoietic cell types was quantified as shown schematically in Figure 3.10a. Figure 3.10b shows poor survival of individually encapsulated UCB cells when the growth factor IL-3 was not added. Furthermore, when bulk exogenous IL-3 was added to the culture, weak enhancement of UCB cell survival was observed, likely due to the limited IL-3 diffusion across the agarose microgel matrix to the UCB cells. In the presence of IL-3 secreting MBA2 cells, a dramatic increase in UCB cell survival was observed, indicating that the local (in- microgel) secretion of IL-3 enabled survival and detection of responsive phenotypes from a heterogeneous input cell population. Importantly, the efficiency of the rescue of the UCB cells varied among the hematopoietic cell types that were present in the heterogeneous culture. Figure 3.10b illustrates that monocytes (CD14+) were most impacted by the co- culture, achieving >80 % rescue, which suggested the importance of IL-3 for the viability of this cell type. Intermediate IL-3 dependent viability was observed in the hematopoietic progenitors (CD34+, CD133+), erythroid cells (GyA+), and granulocytes (CD11b+).

Minimal rescue effect on viability was observed in the megakaryocyte population (CD41+), indicating that IL-3 did not play a role in the survival of this cell type under these conditions.

These results show the differential effect of IL-3 among cell types within a hematopoietic culture system and provide a high-throughput means of screening signalling effects in complex cellular systems. Our experiments also suggest that there are qualitative differences in biological response seen by the local delivery of a soluble factor versus global

(exogenously added) soluble factor delivery, in part due to higher effective concentrations achievable through continuous local delivery.

Given the complex heterogeneity of hematopoietic cell cultures, it has been difficult to discern the varying effects of individual growth factors on specific cell types within the heterogeneous culture. Paracrine signalling cascades and culture dynamics further complicate traditional methods of analysis. The co-encapsulation of the MBA2 cells and the UCB cells enables the determination of the hematopoietic cell types that can be rescued by the localized presence of IL-3. Thus, the MF platform provides a strategy for intricate co-culture studies, while minimizing the complex paracrine interactions that are characteristic for bulk cultures and eliminating difficulties in isolating individual cell types from a heterogeneous culture. In this manner, specific ligand-cell interactions can be interrogated and new biological hypotheses can be developed and investigated.

Figure 3.10 Co-encapsulation of MBA2 cells with UCB cells were used to determine the survival effect of IL-3 on sub-populations within a hematopoietic culture system. (a) Schematic of co-culture of MBA2 cells and UCB cells. (i) Encapsulation of solely UCB cells without exogenous IL-3 results in poor cell rescue. (ii) Addition of exogenous IL-3 (10 ng/mL) results in low rescue of several UCB phenotypes. (iii) Co-encapsulation of MBA2 cells and UCB cells results in low, moderate, and high rescue of specific UCB phenotypes. (b) % Rescue of specific lineages of hematopoietic UCB cells co-encapsulated with MBA2 cells. % rescue of specific UCB phenotypes were measured 3 days after encapsulation in a non-supportive media using eq. 1. This is a representative experiment conducted from at least two independent experiments (n = 2).

3.6 Pre-Microfluidic Experiment Considerations

Over the course of this project, we have encountered several problems which have required modification in the experimental protocol. These problems included wetting on the surface of the

MF device due to cellular release of surface active species, cell death after encapsulation, and the appearance of a bimodal distribution of microgels as a result of the sudden change in shear forces acting on the agarose droplets.

3.6.1 Wetting of the Microfluidic Channel A challenge in MF cell encapsulation is the time-dependent change in the surface properties of the microchannel walls. It has been observed by our group, as well as other research groups, that the wetting properties of PDMS microchannels change during the course of a

cell encapsulation experiment, presumably, due to the release, by cells, of surface-active species such as proteins, carbohydrates, or cell debris. The deposition of these molecules on microchannel walls often leads to their wetting with the aqeous droplet phase and results in poorly controlled droplet generation. This problem can be overcome by the chemical modification of the PDMS channels prior to the encapsulation experiments.

Hydrophobization of PDMS using silanization techniques can be used to prolong the lifetime of PDMS devices when used for cell encapsulation.26

3.6.2 Maintaining Cell Viability

It was observed that encapsulated R1 mES cells had low viability after encapsulation. We hypothesized that the low viability is a result of a single experimental variable or a combination of variables. We performed experiments at varying temperature of the continuous phase in the outlet tubing in the range from 2 oC to 8 oC. These experiments did not show significant improvement in cell viability. The surfactant Span 80 was replaced with the water-soluble surfactant Pluronic F68, a commonly used non-toxic surfactant in cell encapsulation experiments. The change in surfactants did not lead to an increase in viability. We then attempted to use YC5-NEO cells, an mES cell line instead of the R1 cells. Using the original MF encapsulation protocol, the cell viability of the YC5-NEO cells in agarose microgels was greater than 80 %. We hypothesized that the low viability of R1 mES cells (in comparison to YC5-NEO mES cells) was a result of the inherent vulnerability of R1 mES cells to shear stresses imposed by the encapsulation procedure, since other variables (temperature, surfactant, and addition of surface dyes) did not affect the viability of the R1 mES cells in control experiments.

3.6.3 Bimodal Distribution of Particle Size

Preceding the determination of the optimal droplet size (~110 µm) for achieving an encapsulation efficiency of 100% with a cell concentration of 8x106 cell/mL, it was realized that microgels with a bimodal distribution of sizes were formed. During the development of this MF system, the outlet tubing had an inner diameter of 1.23 mm. The diameter of the downstream channel in the MF device had a diameter of 0.15 mm; therefore, a droplet flowing through the main channel of the device experienced a dramatic reduction in pressure as it left the main channel and entered a significantly wider outlet tubing. As a result of the change in pressure, the droplet is broken into two asymmetric droplets, resulting in a bimodal distribution of microgel sizes (Figure 3.11). The sudden change in pressure and direction resulted in the formation of a second ‘T-junction’, where the droplets broke up into two droplets. To avoid this effect, the outlet tubing was exchanged with tubing with a smaller inner diameter (approximately 500 µm).

The replacement allowed the droplets to maintain their integrity and resulted in a monodisperse size distribution.

Figure 3.11 Optical microscopy image of a bi-modal population of microgels containing two distinct sizes. Scale bar is 100 µm.

3.7 Conclusions

The MF platform provides a fast, efficient, and easy to implement method for the high- throughput generation of cell co-culture libraries which can be analyzed by optical microscopy and flow cytometry. The MF strategy offers the speed and the control over the total number of encapsulated cells and the relative numbers of different cell populations. Co- encapsulation and co-culture of MBA2 and M07e cells at varying ratios demonstrated the ability of this approach to modulate paracrine signalling amongst cell populations in a well- defined microenvironment. Furthermore, the MF method can be utilized as a tool to investigate the impact of specific growth factors on a heterogeneous cell population. Co- encapsulation of MBA2 cells with UCB cells was used to determine responsiveness of a sub- population of the hematopoietic cell types that are strongly IL-3 dependent.

The MF approach offers two important advantages: 1) the use of isolated microenvironments provides the capability for studying biological networks, which otherwise cannot be easily interrogated in a bulk environment; 2) The MF platform offers the ability to control the ratio of the two cell populations (a growth factor producing population and a heterogeneous cell population). We demonstrated that the ratio of the encapsulated cells is important as it determines the dose of the signal provided by the secreting cells and received by the receptor cells (and its consequent biological response). These findings indicate that direct co-culture had different biological effects than the addition of a soluble factor.

Further development and improvement of the MF platform will expand the scope of its applications. The ability to screen the phenotypic and functional effects of cell-cell interactions in-situ, ideally with live cells, would prevent the need to remove cells from the beads for analysis. In addition, co-encapsulation of various test cell populations combined

with microfluidically mediated variation in properties of the microenvironment, that is growth factors, mechanical properties (stiffness), or other “niche” parameters would add to the range of conditions that can be studied. Given the rapid development of microfluidic encapsulation technologies, these next steps should emerge shortly.

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Chapter 4 High-throughput Generation of Cell-Laden Microenvironments with Controlled Mechanical Properties 4.1 Introduction

In native tissue, cellular microenvironments have a broad range of mechanical properties. For example, elastic moduli of brain, muscle, osteoid matrix and bone tissues are in the range of 0.1-

1, 8-17, 25-40 and~1.5x107 kPa, respectively.1-3 Currently, it is well established that cells are able to mechanically sense the stiffness of a surrounding matrix by adhering to and pulling upon it using various physio-chemical processes such as the chemical binding of cell surface integrin proteins with various peptide sequences. These processes are followed by a biochemical cascade which activates various processes, such as cytoskeletal cellular response,4 which further dictates the behavior of the cell. The stiffness of the matrix influences many aspects of cell behaviour such as cell adhesion, 5 motility,6 phagocytosis,7 and cellular differentiation.8 Recent, studies of the response of mesenchymal stem cells seeded onto collagen-coated polyacrylamide gels with various elasticities showed that varying the mechanical properties of the substrate resulted in the differentiation of the cells towards specific lineages: the cells were controllably directed towards the differentiation of neurons, myoblasts, and osteoblasts, when cultured on substrates with low, intermediate and high elastic moduli, respectively.9

The stiffness of the matrix has also affected the behavior of cancer cells. It is currently established that the development of cancerous cells is largely determined by genetic mutations and microenvironmental changes, such as the change in the local concentration of a various cytokines as well as changes in local mechanical stimuli.10 Over the past several decades, advances in cellular and genetic interrogation technology have led to greater understanding and

identification of key genetic mutations that are responsible for tumour formation, however, significantly less is known about the role of the mechanical properties of the microenvironment on the behavior of cancerous cells.

The stiffness of the matrix surrounding cells serves an important role in tumorogenesis, as cancerous masses are frequently characterized by their unusually high stiffness within their localized tissue. The increase in the stiffness of the tumour is likely due to the development of the extracellular matrix (ECM) during the process of fibrosis (as a result of tumour expansion), which is the formation of excess fibrous connective tissue. The stiffness of the ECM surrounding cancer cells has been shown to enhance cell motility (metastasis).11 As a result, it has been suggested that an increase in the elastic moduli of the ECM induces cancer cell migration, as it leads to the coordinated migration of cancer cells by stimulating the formation of leader cells and enhancing cell–substrate adhesion. Similarly, in response to mechanical stress, fibroblasts synthesized and secreted different ECM proteins, including fibronectin, tenascin and collagen, and directed matrix remodelling through the expression and secretion of matrix metalloproteinase (MMP) and crosslinking enzymes.12 The ability to degrade and remodel the

ECM allows motile cells to invade local tissue. Current theories have proposed that in contrast with healthy cells, which have the ability to control their self-renewal and proliferation ability based on local cues, cancer cells have lost their ability to sense and respond appropriately to the stiffness of their surroundings matrix. Recently, hepatocellular carcinoma cells encapsulated in a

3D matrix comprised of poly(ethylene glycol) (PEG) hydrogels showed a decrease in malignancy (malignancy was quantified based on angiogenic activity, which was evaluated by measurement of the production of vascular endothelial growth factors (VEGF) and in vivo blood

vessel growth) of the tumor in stiffer matrices, indicating that cancer cell response can be affected by varying the mechanical properties of the surrounding matrix.13

Currently, most of the reported studies on the relationship between the mechanical properties of the microenvironment and cell behaviour are being conducted for cells seeded on planar substrates, yet, in-vivo, cells generally reside in three-dimensional (3D) environments, that is, they are surrounded by a complex and dynamic niche composed of other cells and ECM proteins. Physical forces imposed by the matrix are transmitted to cells in 3D environments in a spatially different way, when compared to cells cultured on planar substrates. Interactions between the cell and the ECM provide physical and biochemical signals that directly affect cell behavior.14 Fabrication of artificial niches has been extensively sought after as a means to accurately control the local cellular environment, and hence, control the behavior of cells.

Encapsulation of cells within 3D hydrogel materials with different stiffness enables the investigation of the effects of the mechanical properties of 3D environments on cell behaviour. A large number of hydrogel matrices (both synthetic and natural) have been used for cell culture15-

18 Many studies have utilized micrometer-sized hydrogels (microgels) to encapsulate cells in a small (<200 µm) microenvironment for cell culture. A size limitation of 200 µm for the microgels allows for easy exchange of nutrients and gases between the encapsulated cells and the surrounding medium.19,20 The preparation of microbeads with varying mechanical properties to study the effect of varying matrix composition on cell fate was recently reported by several groups.13,21,22

Recently, a microfluidic (MFs) exploratory platform have provided a new avenue for fundamental studies in cell biology.23 In particular, MF platforms have been utilized for the high- throughput generation of cell-laden microgels with well-defined properties.24 Encapsulation of 85

cells in droplets or microgels has allowed for studies of cell growth and viability,25-27 gene expression,28,29 or enzymatic activity.30-32 In particular, MF encapsulation of cells in microgel particles offered the following important advantages:

(i) the ability to create 3D cellular microenvironments (niches) with precisely

controlled dimensions and chemical and mechanical properties;

(ii) the ability to vary the properties of microenvironments in a high-throughput

manner;

(iii) the generation of microgels at a rate on the order of tens to hundreds of Hz, which

provided the opportunity to create a large combinatorial library of encapsulated

cells.

In this chapter we report a MF method to high-throughput generation of agarose microgels with varying mechanical properties, which are used for the encapsulation of cells, in order to interrogate the effect of matrix elasticity on metastatic activity. Agarose is a neutral polysaccharide that forms hydrogels at reduced temperatures.33 The mechanical properties of agarose gels can be easily tuned by varying the concentration of the polymer within the microgel.34-37 Agarose was chosen an exemplary matrix material for the study of non-adherent cell behaviour in a confined environment, at varying mechanical stiffness.

We studied the effect of matrix stiffness on the behaviour of non-adherent acute myeloid leukemia-2 (AML2) cells. Acute myeloid leukaemia is a heterogeneous clonal disorder of haemopoietic progenitor cells and the most common malignant myeloid disorder in adults. The

AML2 cells typically reside in the bone marrow before penetrating into the circulatory system.38

Therefore, the determination of the penetration ability at which the cell is able to protrude and escape beyond its native surrounding, at varying stiffness of the microenvironment is highly

desirable, in order to better understand the underlying mechanism of the effect of mechanical properties of the surrounding matrix on cell behaviour.

In this chapter, we report an approach to quantify cell behaviour in microscopic and macroscopic agarose gels. The mechanical properties of agarose microgels were controlled by varying the concentration of the polymer. Cell-laden microgels with varying mechanical properties were produced by MF emulsification of solutions with varying agarose concentration and subsequent fast gelation of droplets. Rapid, throughput change in agarose concentration within the microgels was achieved by supplying in the MF droplet generator two streams of agarose solutions, one solution containing a low concentration of agarose and the other solution contained a high concentration of agarose. By varying the relative volumetric flow rates of these streams, the final concentration of agarose, and hence, the elasticity of the matrix could be tuned. At physiological temperatures (37 oC), the MF method enabled a >10 fold variation of the shear elastic modulus of the agarose microbeads.

4.2 Results and Discussion

4.2.1 Characterization of Macroscopic Agarose Gels

4.2.1.1 Viscosity of Agarose Solutions

To develop a robust approach to the generation of cell-laden agarose microenvironments, we first characterized the properties of macroscopic agarose solutions and gels at varying concentration of agarose. Two important properties are affected by an increase in agarose concentration. Both the stiffness of the gel and the viscosity of the polymer solution increases with increasing concentration of the polymer solution. The second property greatly affects the ability to emulsify the solution using a MF method: at high viscosities, under typical

emulsification conditions, the polymer solution cannot be broken into droplets and instead, forms a thread with nodules, as shown in Figure 1.2 in Chapter 1.

Agarose is a thermally gelling biopolymer which gels upon cooling. At increasing agarose concentration, the viscosity of the polymer solution increases, and the gelation temperature decreases.39 Since microfluidic emulsification of agarose solutions occurred at 37 oC, and we

o measured the viscosity of agarose solutions at varying agarose concentration, Cag, at 37 C, to determine the maximum concentration of agarose solutions that can be readily emulsified in the

MF droplet generator. Figure 4.1 shows the change in viscosity of agarose solution vs. Cag in

PBS buffer. As expected, an increase in Cag led to the increase in viscosity of the solution. The viscosity of agarose solution was ~6 to ~790 cPs for Cag of 1 and 5 wt. %, respectively. A sharp increase in viscosity for the agarose solution was observed for 4 wt. %, in comparison with 3 wt.

%. The increase in the viscosity of agarose solutions at increasing agarose concentration was a result of increasing number of double helices and increasing crosslinking density of the network formed by the double helices.40

Figure 4.1 Variation in the viscosity of agarose solution with increasing concentration of the biopolymer in PBS solution. Measurements were conducted at 37 oC. 1, 2, 3, wt. % solutions were measured with a spindle spin rate of 60 RPM, 4 wt. % was measured at a spindle spin rate of 20 RPM, and 5 wt. % was measured at a spindle spin rate of 8 RPM.

4.2.1.2 Young’s Modulus of Agarose Gels

Concurrently with viscosity measurements, we measured the Young’s moduli (E) of agarose gels with varying Cag by using two independent methods. First, we utilized a conventional compression method to determine the mechanical properties of macroscopic gels.41 We placed a gel disk (10 mm in diameter, 2.3 mm in height) between two plates. The gel disk was loaded onto the microtester and compressed at a rate one-tenth of its height per minute, and held to allow for stress relaxation until equilibrium was reached. The Young’s moduli of the gels were calculated from the ratio of the recorded stress at equilibrium and the applied strain. The compression of the gel was conducted at room temperature.

A second approach to the measurements of the Young’s modulus was performed through the use of an atomic force microscopy (AFM) and compression of the microgels using a tipless cantilever. The cantilever was positioned above the agarose microgel and lowered, until it made contact with the top of the microgel. Measurements were conducted in PBS buffer in the tapping mode. The force required to deform the microgel was measured. The resulting value of E was compared with that attained for the agarose macroscopic gels with the same Cag. Figure 4.2 shows the agreement in the values of E of agarose gels obtained by the two independent methods. Similar to the viscosity measurements, we observed an increase in the Young’s modulus of the microgels with increasing Cag. Figure 4.2 shows the range of E from 0.62 kPa for

1 wt. % agarose microgels up to 20.21 kPa for 5 wt. % agarose microgels, when measured at room temperature.

At 37 oC, the values of E drastically reduced (Figure 4.2) ranging from 0.094 to 4.63 kPa for 2 and 5% agarose, respectively (we were not able to accurately measure the value of E for microgels composed of 1 wt. % agarose). These range of moduli that were achieved share similar mechanical properties to soft tissues found in vivo such as neuronal grey matter, as well as soft muscle and breast tissue.42

4.2.1.3 Encapsulation of AML2 Cells in Macroscopic Gels

Following the characterization of the mechanical properties of the agarose gels at r.t and 37 oC, we prepared cell-laden macroscopic gels with a volume of 20 µL and 1  Cag  5 to determine the effect of matrix stiffness on cell viability. The volumes of the macroscopic gels were

3.0x105 fold larger than that of microgels prepared by the MF method.21

We used AML2 cell line, which is a human leukemia cell line established from peripheral blood of a patient with acute myeloblastic leukemia.43 AML2 cells are non-adherent cells and typically grow in suspension as single round cells. The initial cell concentration in the agarose solution was 1.25x105 cells/mL. Cells were cultured for up to 7 days in cell medium composed of 85%

Dulbecco's Modified Eagle (DME) Medium, supplemented with 15% Fetal Calf Serum (FCS).

The cell-laden gels were analyzed on days 0 and 7. Cell behavior was monitored to follow for cell growth and viability.

Figure 4.3 shows optical microscopy images of AML2 cells embedded in agarose gels at varying

Cag on day 0 (immediately after encapsulation) and on day 7. On day 0, cells were suspended in all agarose gels as suspensions of individual cells. On day 7, distinct differences emerged between the cells embedded in different gels samples. In the gel consisting of Cag = 1 wt. %, the cells showed the greatest growth and the formation of cell colonies (cell clusters). The cells embedded in 2 wt. % agarose showed qualitatively similar results to the cells in 1 wt. % agarose.

The sample containing 3 wt. % agarose showed an increase in the number of cells, compared to day 0, however it was markedly lower, compared to cell growth observed in the samples at Cag of

1 and 2 wt. %. Similar to the cells encapsulated in 1 and 2 wt. %, the majority of cells in 3 wt. % were present in cell colonies, however, the number of cell colonies was fewer and the colonies were smaller in size.

Cells encapsulated in agarose samples at Cag = 4 wt. % showed a minimal growth of cells and the cells appeared as smaller cell colonies then in gels at Cag = 3 wt. %. Very minimal individual cells or cell colonies were observed in the sample at Cag = 5 wt. %.

On day 7 post-encapsulation, fluorescent dyes were added to the cell samples to distinguish between viable and non-viable cells. Hoescht, Calcein AM, and Ethidium homodimer-1 were added to stain for nuclear DNA of all cells (viable and non-viable), viable cells, and non-viable cells, respectively. Figure 4.4 shows exemplary fluorescence imaging of cells immobilized in 1 and 3 wt. % agarose slabs on day 7. As expected, a typical cell colony in the 3 wt. % agarose gel appeared smaller than typical colonies formed in 1 wt. % gel. Furthermore, a larger number of non-viable cells (greater ratio of cells presenting fluorescent emission from Ethidium homodimer-1, as compared to cell presenting fluorescent emission from Calcein AM) were observed at increasing Cag. Cell growth in macroscopic gels was analyzed quantitatively on Day

7. Analysis was performed using a water-soluble tetrazolium (WST) salt. WST-8 in utilized in conjunction with the electron mediator, 1-Methoxy-5-Methylphenazinium Methyl Sulfate (1- methoxy PMS), to assess viable cells. The tetrazolium salt, WST-8, is reduced through a reaction with the reduced form of 1-methoxy PMS. 1-methoxy PMS resides at the cell membrane and reacts directly with NADH (nicotineamido adenine dinucleotide reduced form). NADH is generated from NAD+ by the reaction of dehydrogenase enzymes. Therefore, the tetrazolium salt is utilized for the determination of dehydrogenase activity (Figure 4.5).44 We related the conversion of WST to formazan dye (absorption max at 450 nm) to the number of viable cells in a particular sample.

Optical imaging shown in Figure 4.3 showed a decrease in the number of AML2 cells (on Day 7 compared with Day 0) as Cag was increased. Similarly, on 7 days post-encapsulation of AML2 cells in the macroscopic agarose gels, CCK-8 dye was added to each sample at a 1:10 ratio

(CCK-8 dye:cell culture media). The samples containing the dye were incubated at 37 oC (5%

CO2) for 3 hours. Following the 3 hr incubation, absorbance measurements at 450 nm showed a decrease in the number of metabolically active cells at increasing Cag. Figure 4.6 shows that at increasing Cag, the absorption at 450 nm decreased, due to lower conversion of WST to the

Formazan dye. We attribute the decrease in the conversion of WST to the lower number of metabolically active cells. The graph in Figure 4.6 was normalized relative to the absorbance at

450 nm of the sample composed of cell-laden 5 wt. % agarose (on Day 7), in order to quantitatively compare the relative fold increase in the absorbance at lower Cag < 5 wt. %. Cells encapsulated in gels at Cag of 1 and 2 wt. % agarose showed statistically equal absorbance, with a slightly lower absorbance compared to the agarose-free control (0 wt. % agarose). Both 1 and 2 wt. % agarose samples showed approximately a 3 fold increase in the relative absorbance at 450 nm in comparison with cells encapsulated in gels at Cag = 5 wt. %. From the graph, it is evident that at increasing agarose concentration (> 2 wt. %), the number of metabolically active cells decreases.

Figure 4.5 Structure of WST and its reduction to a Formazan Dye.

Encapsulation of AML2 cells in macroscopic agarose slabs allowed for large-scale quantification of cell behavior, in order to identify the role of the mechanical properties of the agarose gels on cell growth and viability. Based on the results the macroscopic experiments, we utilized the MF system to produce agarose microgels with a variation in Cag from 1 to 5 wt. % (and the variation in Young’s moduli up to 4.63 at 37 oC). The use of MFs technology offers the ability to change

Cag (and hence the mechanical properties of the microgels) in a high-throughput approach by adjusting the relative flow rates of the streams of agarose solutions supplied to the MF device. It should be noted that each stream carries an equal amount of cells.21

4.3 Microfluidic Encapsulation of AML2 Cells

Figure 4.7 shows the microfluidic generation of cell-laden agarose droplets and their subsequent gelation into microgels. Agarose solution carrying cells is injected into the MF device where the liquid stream is subsequently broken into monodisperse droplets due to shear forces exerted by the cross-flow of the continuous phase (mineral oil).

Figure 4.7a shows a schematic of a T-junction MF device which comprises a single channel for the introduction of cell-laden agarose suspension, and second perpendicular channel for the introduction of the continuous phase (mineral oil). Figure 4.7b shows a schematic of a similar

MF geometry, which comprises of two channels for the introduction and mixing of two independent streams of agarose. Following the mixing of the agarose solutions, the agarose solution is emulsified into monodisperse droplets. Mixing of the two liquids prior to emulsification allows for high-throughput variation in the final ratio of the two solutions. We previously reported a MF device which utilized the supply of two streams of agarose solutions with different concentrations, which upon mixing, would be emulsified into monodisperse droplets.21 This approach allowed for the high-throughput variation in the concentration of agarose (and hence the elasticity) of the microgels. A schematic of the MF device for the high- throughput variation in the composition of agarose microgels is shown in Figure 4.7b.

To develop a method for MF generation of cell-laden agarose microgels with varying elasticities

(microenvironment libraries) for studies of AML2 cell behavior, we considered the following requirements:

(i) The concentrations of agarose in the concentrated and dilute solutions (supplied as

Streams 1 and 2, respectively) had to be such that sufficient mixing occurred in the

microchannel prior to emulsification of the mixed stream in the MF droplet generator

(ii) Microgels with different elasticities had to have identical average dimensions.

(iii) The encapsulation efficiency should be governed by the Poisson distribution45, as

described in Chapter 3.

(iv) The ratio of viscosities of the two agarose solutions should be such that no back flow

occurs in the MF device.

Initially, we explored the preparation of cell-laden agarose microgels by using a single precursor solution of cell-laden agarose. We utilized a single channel geometry as an exemplary tool to produce cell-laden microgels with mechanical properties, which are controlled by the value of

Cag of the precursor agarose solution. Figure 4.7a shows a schematic of a MF device comprising a single inlet channel for the disperse phase. As the disperse phase is introduced into the MF device, it travels and enters the T-junction. Droplet formation occurs in a highly periodic breakup regime, due to the shear forces exerted on the disperse phase (a suspension of cells in agarose solution in PBS) by the continuous oil phase (mineral oil).46 In our experiments, the total flow rate of the disperse phase was maintained at 0.1 mL/hr. The flow rate of the continuous phase was tuned, in order to produce droplets with a mean diameter of 110 µm (yielding 100

µm-diameter cell-laden microgels) as discussed in Chapter 3.

Figure 4.7 Schematic of the MF device for the generation of cell-laden agarose microgels with tunable elasticity. (a) MF device comprising one inlet channel for the disperse phase. (b) MF device comprising two inlet channels for injection of the disperse phase. The height of both devices was 150 µm. The width of the horizontal channels supplying the mineral oil phase and the width of the channel at the point of the T-junction was 150 and 20 µm, respectively. The length of the mixing channel prior to the T-junction was 250 mm.

We prepared agarose solutions of AML2 cells suspended in the droplet phase to a concentration

6 of 110 cells/mL. The microgels were produced at Cag = 1 and 2 wt. %, which corresponded to

E  0.62 and 2.56 kPa at room temperature. Following incubation at 37 oC, the 2 wt. % agarose microgels attain an elastic modulus of 0.094 kPa. The microgels had a round shape and an average diameter that was approximately 10% smaller than the mean diameter of the corresponding droplets. The microgels had a narrow size distribution, with a polydispersity that did not exceed 2.5%. We stress that in comparison with our earlier work described in Chapter 3, we aimed to minimize the number of microgels containing more than one cell by reducing the initial concentration of cells in the feed solution.

Figure 4.8 shows the theoretical and experimental results for the percentage of microgels which contain 0, 1, 2, or 3 cells. Theoretical values for the percent of the number of cells per microgel were calculated using the Poisson distribution (described in detail in Chapter 3), based on a droplet diameter of 110 µm, and an initial cell concentration of 1x106 cells/mL. For the precursor droplets with a mean diameter of 110+/- 2.8 m, we achieved an overall encapsulation efficiency of 45.4 +/- 3.9%, with 34.2 +/-5.0 % of the microgels containing one cell, 9.0 +/-

2.6% two cells and 54.6 +/- 3.9% containing no cells (empty). Figure 4.8 shows excellent agreement between experimental and theoretical results attained using the Poisson distribution.

The representative images of AML2 cells encapsulated in agarose microgels at Cag of 1 and 2 wt.

% are shown in Figure 4.9 on Day 0 (immediately after encapsulation and transfer into cell media) and on Day 7 (in cell media). Immediately following the encapsulation of the cells in agarose microgels cell viability was determined to be 64.8 ± 1.2% using a standard trypan blue staining and quantification using a haemocytometer. The relatively low cell viability of AML2 cells needs to be further explored.

100

Figure 4.8 Comparison of the number of encapsulated cells per droplet compared to theoretically expected Poisson distribution.The graph shows the number of cells per microgel determined experimentally (■) and theoretically (for 110 µm-diameter droplets) using the Poisson distribution (●). The concentration of cells in the feed suspension was 1x106 cells/mL. Approximately 300 microgels (formed at Cag = 2 wt. %) were analyzed to determine the encapsulation efficiency. The diameter of the precursor droplets was 110 +/- 2.8µm. QOil = 1.2 mL/hr, Qag = 0.1 mL/hr.

Comparison of optical images of cell-laden microgels on Day 7 with images taken directly after encapsulation, cell growth appeared to be minimal in microgels composed of either 1 or 2 wt. % agarose. The reasons for the limited cell growth are not well understood and remain under investigation.

101

4.4 Conclusions

The results of our work pave the way for the high-throughput generation of 3D cellular microenvironments with varying mechanical properties. The microgels can be used for subsequent cell culture, thereby enabling efficient studies of the effect of mechanical properties on cell fate. The properties of cells can be examined within the microgels using various spectroscopic techniques such as absorption (UV-Vis) and fluorescence microscopy (fluorescent microscopy, confocal microscopy), or after release of the cells from the gel matrix through the digestion of the gel network using enzymatic cleavage (e.g. agarase).

In the present work we achieved a moderate range of the elastic moduli of agarose hydrogels at r.t. and at physiological temperature (37 oC). Future work will involve the generation of cell- 102

laden microgels with increasing Young’s modulus (up to give the value), in order to investigate further the local effects of the mechanical properties of the microenvironment on cell behaviour.

Currently, we achieved the encapsulation of AML-2 cells in 1 and 2 wt. % agarose microgels.

Further optimization of cell viability will need to be addressed in order to ensure successful and accurate sample preparation and analysis of cell behaviour.

In addition, the exploration of other biopolymers (e.g. collagen, fibronectin, elastin, etc.) offers interesting and exciting opportunities for studying the effect of both the mechanical and chemical microenvironments of cells. In addition, the use of two inlet channels for two independent disperse phase solutions will allow for the high-throughput generation of agarose microgels with varying elastic properties. Further measurements of the elastic moduli of the microbeads will be optimized using atomic force microscopy.47 These experiments will be extended to microbeads compartmentalizing cells in the course of cell culture, and the correlation between the varying elasticity of the microbeads and cell behavior.

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(26) E. Brouzes, M. Medkova, N. Savenelli, D. Marran, M. Twardowski, J. B. Hutchison, J. M. Rothberg, D. R. Link, N. Perrimon, M. L. Samuels, Proc. Natl. Acad. Sci. U. S. A., 2009, 106, 14195.

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(29) A. Huebner, M. Srisa-Art, D. Holt, C. Abell, F. Hollfelder, A. J. Demello, J. B. Edel, Chem. Commun., 2007, 1218.

(30) A. Huebner, L. F. Olguin, D. Bratton, G. Whyte, W. T. S. Huck, A. J. de Mello, J. B. Edel, C. Abell, F. Hollfelder, Anal. Chem., 2008, 80, 3890.

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(33) A. Hayashi, T. Kanzaki, Food Hydrocolloids, 1987, 1, 317.

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(37) S. Lahooti, M. V. Sefton, Cell Transplant., 2000, 9, 785.

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Chapter 5 Integrated Microfluidic Reactors for Studies of Chemical

and Physical Processes

5.1 Characterization of On-Chip Reactions and Integrated Analytical Tools

A wide range of chemical processes have been performed on-chip using both single phase and two phase MFs. These reactions vary from on-chip organic reactions1-3 to the synthesis of nanoparticles,5-7 and protein crystallization.8,9 Characterization of the products and the transient species of a chemical reaction during the reaction process is paramount in the optimization of reaction conditions. In conventional reactions, small samples are periodically removed from the reaction chamber and analyzed to monitor the state of the reaction. Microfluidic optimization of reaction conditions requires the same periodic analysis of the reaction, however, currently the majority of on-chip characterization is performed via optical microscopy or fluorescence-based optical microscopy.10 Several groups integrated analytical tools for direct on-chip characterization. Among these are on-chip NMR,11,12 fluorescence,10 infrared (IR),13,14 Raman spectroscopy,15 surface enhanced Raman spectroscopy (SERS),16 and pH probes.17,18 Yet, in many instances, the reactions can be monitored more accurately (and efficiently) when more than one analytical tool is used. For instance, organic reactions which utilize mineral acids (e.g. SN1 reaction between t-butanol and HCl) can be monitored by real-time ATR-FTIR spectroscopy. In this process, ATR-FTIR can only monitor the changes in vibrational bands of the organic reactant as it transforms into an organic product, but cannot monitor the change in the depletion of the mineral acid (HCl). Simultaneous characterization using an in-situ pH probe would

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provide direct real-time information about the concentration of the acid during the reaction. An example of a multi-probe system was recently reported as a platform for on-chip cell culture. The

MF system incorporated several analytical probes for continuous characterization of cellular conditions.19 Other systems have also incorporated multiple analytical tools for direct on-chip characterization.20,21 The device included a pH probe, a temperature probe, and a dissolved oxygen sensor. Feedback from the three probes provided continuous computer controlled adjustments to the media conditions. This system demonstrated the capability of a fully integrated MF platform. Future improvements in this system will include interchangeable probes, such that one could change or replace one probe with another depending on the requirements of the experiment.

5.2 Microfluidic Studies of Carbon Dioxide Dissolution

Typical methods used to interrogate gas-liquid reactions have a large laboratory foot print, are complex, and require long acquisition times to collect sufficient data.22-24 For example, one method utilized a high pressure chamber filled with an aqueous mixture of methyldiethanolamine

25 and diisopropanolamine. A supply of CO2 gas into the chamber allowed for the formation of

CO2 bubbles. Carbon dioxide bubbles were generated until the solution reached a gas/liquid (the experiments were repeated at various temperatures). In some instances, two weeks were required to achieve equilibrium (depending on the composition of the aqueous solution and the temperature of the reaction). Further difficulties with these methods can arise due to a lack of control over the delivery of the gas, as well as inconsistency in characterization of the sample due to a lack of integrated analytical tools; in many cases, the sample is removed from the pressure chamber for analysis.

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In comparison with most macroscopic systems which are limited to studies of gas-liquid reactions at equilibrium, MF based systems can study the evolution of a reaction from the beginning until the reaction reaches equilibrium. This feature enables the acquisition of kinetic data in a high-throughput manner in a short period of time with minimal use of reagents.

Recently, several groups demonstrated MF routes to high-throughput generation of CO2 plugs in various industry standard CO2 sequestering solvents. The MF platforms provided an accurate control over the supply of the gas and the liquid phases. The change in plug size was monitored vs. time using a high speed camera. The change in bubble size was used to relate the extent of mass transfer of the gas into the liquid slugs. Currently, MF studies have been focused on the

28,30-27 28 29 dissolution of CO2 in different solvents at varying temperature and pH. Recently,

Abolhassani et al. developed an integrated computer controlled high-speed data acquisition MF system capable of characterizing the extent of dissolution of a gas (by measuring the change in bubble size) in a particular solvent within a very short time scale (minutes),26 which favorably differs with macroscopic experiments, which would require days to weeks to compile a comparable data set. As MF technologies improve, the ability to change and control reaction conditions such as temperature,28 pH,29 and molar ratios of reagents,30 will allow for a broader study of reactions at varying conditions.

5.3 Summary

Microfluidic technology offers an exciting avenue for studying chemical reactions with a high degree of control over reaction conditions with minimal reagent consumption. The integration of probes to monitor in-situ reaction conditions in real-time offers an extended asset as it allows for direct characterization of the reaction with various analytical techniques in parallel. In this regard, a highly robust and accurate system for in-depth investigations of complex gas-liquid

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systems should yield important information about the fundamental interactions which are difficult to acquire using conventional bulk setups. Among the many advantages of MF platforms, segmented flow has provided a means for high-throughput generation of bubbles for studies of gas-liquid reactions. Such systems will be beneficial for future studies of many important greenhouse gases such as CO2, CH4, NOx, and SO2. The ability to accurately characterize the early evolution of ultra-fast reactions will be useful for chemist, biochemists, and engineers.

References

(1) Z. T. Cygan, J. T. Cabral, K. L. Beers, E. J. Amis, Langmuir 2005, 21, 3629.

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(10) G. Ryu, J. Huang, O. Hofmann, C. A. Walshe, J. Y. Y. Sze, G. D. McClean, A. Mosley, S. J. Rattle, J. C. deMello, A. J. deMello and D. D. C. Bradley, Lab Chip, 2011, 11, 1664.

(11) J. Bart, A. J. Kolkman, A. J. Oosthoek-de Vries, K. Koch, P. J. Nieuwland, H. Janssen, P. J. M. van Bentum, K. A. M. Ampt, F. Rutjes, S. S. Wijmenga, H. Gardeniers and A. P. M. Kentgens, J. Am. Chem. Soc., 2009, 131, 5014.

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Chapter 6 Temperature-Controlled 'Breathing' of Carbon Dioxide

Bubbles

6.1 Introduction

Since the start of the industrial age, the increase in levels of atmospheric carbon dioxide

(CO2) have had a negative environmental impact due to contributions to global warming, ocean acidification, and related abnormal change in marine ecology.1 These trends have sparked intense interest in the chemical and physical behaviour of CO2 with the environment.

Currently, studies of CO2 transfer between the atmosphere and the oceans have led to the development of both chemical and physical models for CO2 sequestration and storage, and became a frontier area of collaborative research in chemistry, biology, geology, environmental sciences and chemical engineering.

An important property of CO2 is its temperature-dependent solubility in liquid media: the

2-4 solubility of CO2 in water and organic solvents decreases with increasing temperature. One of the important consequences of this effect is the temperature-dependent transport of CO2 between the atmosphere and the ocean (the so-called “solubility pump”).5,6 At high latitudes the gas dissolves in cold ocean water and at equatorial latitudes it is released from warm ocean waters to the atmosphere. Rising global average temperatures have been linked to decreasing solubility of CO2 in the oceans. Since the oceans are the largest sink for atmospheric CO2, this effect may have dramatic consequences on the ability of seawater to

1 uptake CO2. Currently, the magnitude of potential feedback mechanisms on CO2 levels in

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the atmosphere due to changing temperature and acidity of the oceans and the change in effectiveness of the solubility pump are being addressed primarily via modelling.7 Field studies of CO2 dissolution in ocean water are expensive and time-consuming. An accurate study of a particular chemical or physical effect of CO2 with the environment is challenging, owing to the large number of variables naturally present in nature.

Temperature-controlled CO2 solubility is also important in CO2 sequestration. For subsurface geologic storage of CO2, the ability to delineate the relative importance of solubility, and ionic, and mineral trapping is dependent on a detailed understanding of the interrelated

8,9 effects of temperature, salinity and pH on CO2 solubility. The reversible absorption of CO2 by organic solvents, e.g., perfluorcarbons10 or dimethyl ethers of poly(ethylene glycol)3,11 is another method to remove CO2 from industrial flue gas streams. Regeneration of the solvent is achieved by heating the solvent, thereby reducing CO2 solubility and releasing gaseous

CO2. Furthermore, chemical reactions driven by chemical binding of CO2 to organic solvents, e.g., secondary amines, offer the ability to generate a “solvent polarity switch”: the solvent becomes polar under binding of CO2 but switches back to low molecular polarity

12-14 when CO2 is removed by heating. The solvent polarity switch has a promising green chemistry application by eliminating the need to replace solvents in multi-step processes, which require different solvent properties in different stages of a reaction.

In order to understand the role of various factors on temperature-dependent CO2-liquid interactions, it is important to study them in a controlled manner and on a short time scale

(sometimes, under conditions that are far from equilibrium). The latter requirement is especially important for kinetically controlled processes. The experiments require (i) a controlled supply of CO2 gas to the liquid medium, (ii) the ability to precisely control the

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temperature of the liquid, and (iii) an accurate method of characterising the amount of CO 2 uptaken or released by the liquid. Currently, studies of the dissolution of CO 2 are performed by generating bubbles of CO2 in macroscopic liquid-filled vessels and monitoring the change in bubble size.15 This method lacks control over the size of the generated bubbles, which leads to uncertainties in studies of the kinetics of mass transfer across the gas-liquid interface. Alternatively, injection of liquid CO2 into a vessel containing a large volume of a liquid is followed by studies of CO2 uptake and monitoring the change in pressure inside the vessel.16,17 This time-consuming method requires a complicated experimental setup and high pressure applied to CO2.

In this chapter, we report a microfluidic (MF) platform for studies of temperature mediated

CO2 transfer between the gas and the liquid phases on short time scales. Micro-meter diameter CO2 bubbles with a narrow size distribution were generated in a liquid continuous phase and subjected to temperature modulations as they travelled through the MF channels.

In response to a cooling-heating-cooling cycle, the bubbles exhibited contraction-expansion- contraction transitions, which originated from the temperature-dependent CO2 solubility. Our

MF strategy satisfied the three crucial aspects described above: (i) a controlled supply of

CO2 to the aqueous phase was achieved by the generation of CO2 bubbles with a narrow polydispersity;18-21 (ii) the ability to achieve site-specific control of temperature in the MF device, and (iii) on-line monitoring of mass transfer between the gaseous and the liquid continuous phases, which was achieved by measuring the temperature-dependent change in bubble size.

The MF strategy was demonstrated for the temperature-controlled sequestration of CO2 by four exemplary systems: a dimethyl ether of poly(ethylene glycol), a solvent that does not chemically

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react with CO2 (that is, a physically absorbing environment), deionized water, a 0.7 M aqueous solution of NaCl, and ocean water extracted from Bermuda coastal waters. Analysis of the changes in bubble dimensions allowed an estimation of the effect that temperature and salinity have on the solubility of CO2. The proposed method offered a promising operational platform for kinetic (non-equilibrium) studies of the effect of temperature on CO2 sequestration.

Finally, a similar approach was adapted for the temperature mediated control over the dissolution of CO2 bubbles for the controlled generation of armored bubbles. Bubbles coated with a dense layer of colloid particles (armoured bubbles) show high stability against coalescence,22,23 reduced rate of release of the entrapped gas,24,25 and the ability to acquire anisotropic shapes.26 Armoured bubbles show potential applications in the fabrication of thermal and acoustic insulators, lightweight materials and electrets.27 Conventional methods of the generation of particle-coated bubbles include bulk emulsification of a gas,23,28,29 injection of bubbles in a suspension of particles,30 or flow-assisted assembly of colloids on the surface of bubbles.31 These methods either possess low productivity,31 or generate bubbles with an insufficient control of bubble dimensions.30 Recently, our group reported a chemically mediated MF strategy for producing bubbles coated with micrometer-size particles,32 biopolymers,33 and nanoparticles.34 In particular, bubbles encapsulated with a colloidal armour shell were generated by three concurrent processes occuring within 10 sec: (i) continuous MF generation of CO2 bubbles in an aqueous dispersion of anionic (carboxylated) colloidal particles,

(ii) the dissolution of CO2 leading to the local reduction in pH in the vicinity of the bubbles, and

(iii) pH-mediated protonation of the carboxylic groups on the surface of the microbeads and assembly of hydrophobized particles on the bubble surface. Control over the size of armoured bubbles was achieved by varying the value of pH of the continuous aqueous phase: armoured

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bubbles with smaller dimensions formed under basic (up to pH = 14) conditions, owing to

32 increased efficiency of CO2 dissolution. This approach, however, is not applicable to particles which are unstable under strongly basic conditions, e.g., SiO2 and TiO2 nanoparticles. In our work, we utilized the temperature dependent solubility of CO2 to generate armoured bubbles.

6.2 Background

4 The physical absorption of the gaseous CO2 by the liquid phase follows Henry’s law. In an aqueous environment, physical mass transfer of CO2 is accompanied by chemical reactions with water. These two processes are described by equations 1-3:35

PCO2 = kH[CO2]g (1)

2- + pH > 10: CO2(aq) + H2O ↔ CO3 + 2H K (2)

- + pH < 10: CO2(aq) + H2O↔HCO3 + H K1 (3)

where kH is the temperature-dependent Henry’s law constant, PCO2 is the partial pressure of the CO2 gas, and K and K1 are the equilibrium constants. The relationship between the gas solubility and temperature is reflected by the temperature dependence of Henry’s constant.

3 3 -1 For example, Henry’s constants for CO2 in water are 1.3710 and 3.4510 kPa L mol at

1 and 30 oC, respectively.4 The equilibrium constants are also temperature-dependent,

-7 however their contribution to the dissolution of CO2 in water is weaker (K1 = 2.6910 and

-7 o 4 4.7910 at 0 and 30 C, respectively). We note that the reaction of CO2 (aq) with water is pH-dependent (reactions 2 and 3). In the present work, the original value of pH of the continuous aqueous phase did not exceed 7.95, hence the uptake of CO2 was reflected by equations 1 and 3.

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Figure 6.1 illustrates solubility-dependent shrinkage and expansion of CO2 bubbles: bubbles shrink upon cooling, due to the increased solubility of CO2 in the liquid phase, and expand when the temperature increases, owing to the reversed gas transfer from the liquid to the gaseous phase. This “breathing” of the CO2 bubbles originates from the two-way mass transfer between the gaseous and the liquid phases, controlled by the temperature of the liquid. We assumed that the temperature-dependent solubility of CO2 can be monitored by measuring the change in dimensions of CO2 bubbles when they are subjected to the variations in temperature of the continuous liquid phase.

6.3 Experimental Design

Figure 6.2a shows a schematic of the MF device, fabricated in polydimethylsiloxane

(PDMS), which was used for studies of temperature-dependent dissolution of CO2 bubbles.

The CO2 gas was supplied to the central channel under pressure PCO2=48.3 kPa or PCO2=82.7 kPa through a polytetrafluoroethylene tubing, attached to a Bellofram pressure regulator. A continuous liquid phase at a temperature of 23 +/-1 oC was introduced into the two side channels at the flow rate Qc = 2.5 mL/hr, using a syringe pump. The inset in Figure 6.2a illustrates the generation of CO2 bubbles as the gaseous stream is focused in the narrow

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orifice by the shear force imposed by the continuous phase. A highly periodic break-up of the gaseous thread yielded bubbles with a narrow size distribution.18 The bubbles moved through a serpentine downstream channel with a length of 450 mm. Figure 6.2b shows the variation in temperature along the MF device, plotted as a function of the distance from the orifice

(open squares). Due to the spatial proximity of Regions 1, 2 and 3, a gradient in temperature was established. The temperature of the continuous aqueous phase encountering the gas stream was 11+/-1 oC at the orifice. The lowest temperatures in Regions 1 and 3 were 1 and 3 oC, respectively, achieved at the distances of ~50 and ~450 mm from the orifice, respectively. The highest temperature in Region 2 was 31oC, achieved at a distance of ~300 mm from the orifice. (The temperature range of 1 to 31 oC was selected to reflect the average variation in temperature of the surface of the ocean in the polar and equatorial regions of the Earth, respectively).36 A slightly higher minimum temperature of the aqueous phase achieved in Region 3 (vs. Region 1) originated from the higher temperature of the liquid entering Region 3 from Region 2, in comparison with the initial temperature of 11 oC of the liquid entering Region 1. Figure 6.2b (dotted line) shows simulation results of the temperature of the liquid travelling through the MF device. The results of simulations were in agreement with experimentally determined temperatures. We attribute a small deviation between the simulated and experimental results in Region 2 to a greater than expected heat loss, compared to the simulation parameters, which accounted for ideal heat transfer.

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Figure 6.3 shows a schematic of the experimental setup with distinct temperature zones.

Regions 1 and 3 of the channel were cooled by placing underneath them a hollow copper block and purging through it a glycerol/water mixture (1:4 v/v) at a temperature of 0 oC. The temperature of the mixture was controlled using a water circulator. Region 2 of the microchannel was heated by placing beneath it a heating module with a top surface area of

1.5 cm2. The temperature in the module was controlled using an electronic temperature controller. The temperature of the cooling block and the heating elements were set to 0 and

35 oC, respectively, and the system was equilibrated for 30 min. The temperature of the continuous liquid phase along the MF device was measured using a thermocouple.

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Experiments were conducted in dimethyl ether of poly(ethylene glycol), deionized water, a

0.7 M aqueous solution of NaCl, and ocean water (later in the chapter, the last three liquids are referred to as “water”, “salt water” and “ocean water,” respectively).

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6.4 Results and Discussion

6.4.1 Bubble Breathing in Water

Figure 6.4 shows representative optical microscopy images of the CO2 bubbles dispersed in

deionized water at varying temperatures. The bubbles generated at 11 oC rapidly and

dramatically shrank in the cooled Region 1, expanded in the heated Region 2 and

subsequently, shrank in the cooled Region 3. The entire process - from the generation of bubbles to the moment when they exited from the MF device - occurred within ~19.5 s.

Figure 6.5a shows the variation in the volume of CO2 bubbles in water, plotted as a function of the distance from the orifice (~300 mm). The variation of the experimentally determined temperature in the MF device is shown on the same graph. The shrinkage of the bubbles

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started immediately after the orifice and continued up to a distance of 100 mm from the orifice. At Point 1 (1 oC) the bubbles experienced a 24.4 fold loss in volume. Subsequent heating to 31 oC (Point 2) led to a ~4.4-fold increase in the average bubble volume, in comparison with that at Point 1. Cooling of the liquid to 3 oC resulted in a ~3.2-fold contraction (Point 3), in bubble volume in comparison with that at Point 2.

Figure 6.5b shows the temperature-controlled cycle in the variation of bubble volume. The cycle started at Point 1 and was completed at Point 3. The cooling-heating-cooling events led to a contraction-expansion-contraction cycle of the CO2 bubbles, which we term 'bubble breathing'.

Whereas Figure 6.5 presents a dynamic, non-equilibrium response of bubbles to the changes in temperature. Figure 6.5c shows a control experiment performed at constant temperature. In this experiment, we verified that the volume of the bubbles stabilized at a distance of 300 mm from the orifice, which suggested that at this point, the transfer of CO2 from the gaseous to the aqueous phase has reached an equilibrium.21

The shrinkage of the bubbles was caused by the dissolution of CO2 and its chemical reaction with water described by eqs.1 and 3. The trend in the variation in bubble volume upon cooling and heating followed the change in Henry’s constant (which is ~2.5 greater at 1 oC than at 30 oC),4 thereby compensating for the change in equilibrium constant in eq. 3.

We note in Figure 6.5a a lag in the variation in bubble volume in response to the change in temperatures. We attribute the temporal delay in the change in bubble volume to the time required for the CO2 molecules to diffuse towards or away from the bubble. The lag in bubble contraction and expansion in Regions 1 and 2 occurred over a distance of ~50 mm,

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corresponding to a time interval of ~1.7 s. For the length scale on the order of the bubble

2 size, we calculate the diffusion time as t~D /DCO2, where D is a representative diffusion length, which we define as the average distance from the wall of the channel to the surface of the bubble. To determine the average diffusion length, we averaged the length from the surface of the largest (~100 µm) and smallest (~40 µm) bubbles to the channel walls. The

3 2 -1 37 diffusion coefficient (DCO2) is of CO2 in water is, DCO2 ≈ 1.94 x 10 µm s . For deionized

water, the parameters were: PCO2 = 48.3 kPa, Qc = 2.5 mL/hr, and an average diffusion length of 65 µm (calculated as the average of the half channel width minus the radius of the bubble), the estimated diffusion time of ~2.2 s was similar to the experimentally observed lag time of ~1.7 s.

6.4.2 Bubble Breathing in Salt Water and Ocean Water

Figure 6.6a shows the variation in the temperature-controlled volume of CO2 bubbles in ocean water and salt water. The bubbles exhibited a contraction-expansion-contraction behaviour in both liquids. The volumes of the CO2 bubbles in the salt water and ocean water

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were identical in Regions 1-3 of the MF device. At Point 1 at 1 oC, the volume of bubbles decreased by ~ 17.9 fold, in comparison with their initial volume. At Point 2 at 31 oC, the volume of bubbles featured a ~2.3 fold increase, in comparison with that attained at Point 1.

A~1.9 fold shrinkage occurred at Point 3 (3 oC), in comparison to the volume attained at

Point 2.

Figure 6.6b shows measured values of pH of the ocean water at Points 1, 2 and 3. The variation in the acidity of the continuous phase correlated with the contraction-expansion- contraction behaviour in bubble volume. At temperatures of 1 and 3 oC (Points 1 and 3, respectively) the values of pH were 4.83 and 4.80, respectively; at Point 2 (31 oC), we measured pH = 4.92.

We ascribe a weaker 'bubble breathing' effect in ocean and salt water, than in water, to the non-ideal properties of these liquids. In aqueous electrolyte solutions Henry’s constant is 4

KHo = KH∙γo (4)

where KHo is Henry’s constant in deionized water, KH is Henry’s constant at a particular ionic strength of the medium, and γo is the activity coefficient (γo<1). For comparison, at 30 o 3 3 -1 C, KHo=3.4510 and KH = 510 kPa L mol , for CO2 in deionized water and ocean water, respectively.4 The variation in pH (Figure 6.6b) can be explained by applying Le Chatelier’s principle. As shown in eq. 3, the reaction of CO2(aq) with water generates ions. At lower temperatures, a stronger dissolution of the bubbles resulted in a higher amount of CO 2(aq) reacting with water, which led to an increase in the concentration of H+ ions. Similarly, at a higher temperature, bubble expansion resulted in the removal of CO2(aq) from the continuous phase. By applying Le Chatelier’s principle, we conclude that the removal of CO 2

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from the liquid phase resulted in the reverse reaction, thereby decreasing the concentration of

H+ in the continuous phase, and leading to an increase in the pH of the continuous phase.At increased temperatures, in comparison with water, the presence of cations in salt and ocean water suppressed the transfer of CO2 from the liquid phase into the gas. We ascribe this

40 effect to the reduced vapour pressure of CO2 under non-ideal conditions. The binding

+ energy of dissolved CO2 molecules to Na ions (the most abundant cations in the ocean

38 water) is ~12.7 kcal/mol (vs. ~2.89 kcal/mol for the binding of aqueous CO2 molecules to water39).

We note that due to the presence of Ca2+ ions in ocean water, the formation of non-soluble

40 carbonate salts such as CaCO3 could affect the solubility of CO2. Figure 6.6a shows that a similar size of CO2 bubbles (measured at the same temperatures) in salt water and ocean water suggested that for this range of water activity the binding of carbonate and bicarbonate

2+ ions to Ca ions did not affect CO2 solubility. 127

6.4.3 Bubble Breathing in Dimethyl Ether of Poly(ethylene glycol)

We examined temperature-controlled variation in dimensions of the CO2 bubbles dispersed in dimethyl ether of poly(ethylene glycol) (DEPG), a solvent utilized for removal of acidic gases from industrial exhaust.3,11 Figure 5.7a shows the variation in the bubble dimensions, along with characteristic images of bubbles taken at 1, 31, and 3 oC (Points 1-3). Figure 6.7a shows that similar to the aqueous systems, the contraction-expansion-contraction behavior

o was observed for CO2 bubbles in DEPG. At the point of maximum dissolution (at T = 1 C), we observed 98.9 % shrinkage in volume. We defined a unit volume as a volume of the

22 liquid containing a single initial CO2 bubble (imaged immediately after the orifice). By applying the same unit volume (= 3.8x10-9 L) to the position of the bubble in the downstream channel, we found the relative change in bubble volume as VT/Vo where V0 and VT are the initial and the temperature-dependent bubble volume, respectively. In this manner, we determined the variation in the amount of CO2 dissolved per unit volume at various temperatures as shown in Figure 6.7b. The concentration of dissolved CO2 reduced from

1.79x10-2 to 1.69x10-2 mol/L, when the temperature increased from 1 to 31 oC.

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Figure 6.7 (a) Variation in the volume of CO2 bubbles in DEPG and in the temperature of DEPG, plotted as a function of the distance from the orifice of the MF device. The lines are given for eye guidance. Images (from left to right) show bubbles at temperatures of 1, 31, and 3 o C, respectively. (b) Variation in the amount of dissolved CO2, plotted as a function of the experimentally determined (black symbols) and simulated (white symbols) temperatures. (c) Variation in bubble volume in DEPG, plotted as a function of the distance from the orifice at 23 o C. PCO2 = 82.7 kPa, Qc = 2.5 mL/hr. The scale bar is 50 µm.

6.4.4 Generation of Armoured Bubbles

The effect of temperature on the solubility of CO2 in liquids can be utilized for controlling the extent of dissolution of bubbles in a liquid media under specific conditions. By using MF emulsification, we generated CO2 bubbles in an aqueous dispersion of anionic colloid particles and subjected the bubbles flowing along microchannels to various temperatures. The temperature-controlled dissolution of CO2 led to the generation of armoured bubbles with well- defined reduced dimensions. This method offered an efficient complementary strategy to pH- mediated control over bubble dimensions,21 owing to the ease of tuning the dimensions of armoured bubbles in-situ and the applicability of the method to a broad range of colloidal particles that are unstable under high pH values.

The dissolution of CO2 in water is described by eqn. 1-3, where eqn. 2 can be rewritten in the

- 35 form of the dissociation reaction of HCO3 in water at pH > 10:

K2 - 2- + pH > 10 HCO3 ↔ CO3 + H (5)

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-11 -11 o The values of the equilibrium constant K2 are 2.37x10 and 5.13x10 at 0 and 30 C,

4 respectively. Because of the temperature-dependent solubility of CO2, it is expected that the bubbles will shrink upon cooling, due to the enhanced CO2 transfer from the gas phase to the liquid phase, and expand upon heating, owing to the reversed gas transfer. As we have shown earlier in this chapter, the variation in temperature of the aqueous continuous phase enabled control over the dissolution of CO2 bubbles. We therefore hypothesized that a similar approach can be used for controlling the dimensions of armoured bubbles.

The response of the CO2 bubbles to the variation in temperature was examined in the MF device shown schematically in Figure 6.2a. The CO2 bubbles were produced in the MF flow-focusing bubble generator in a particle-free aqueous continuous phase at 23 oC.18 In the downstream channel, the bubbles travelled through three regions, where they were subjected to the reduced temperature (Regions 1 and 3) or the elevated temperature (Region 2). The corresponding expected change in bubble size is shown schematically next to each zone.

The CO2 gas was supplied to the central channel at a pressure of 20.7 kPa. An aqueous 2 wt. % solution a non-ionic surfactant Triton 100 at pH = 6.0 was supplied to the two side channels at a flow rate of 3.0 mL/hr. The CO2 bubbles were formed by a periodic breakup of the gaseous stream, due to the shear forces imposed by the liquid phase on the gas stream.18 The bubbles travelled through the three regions in the downstream channel (Regions 1-3 in Figure 6.2a), in which the temperature varied from 1 oC to 77 oC.

Figure 6.8a shows optical microscopy images of the CO2 bubbles subjected to various temperatures. The bubbles exhibited a contraction-expansion-contraction behavior, which was consistent with the temperature-mediated solubility of CO2: at lower temperature, the solubility of CO2 in the continuous phase increased, leading to a reduction in the dimension of the bubbles,

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and at elevated temperatures the solubility of the CO2 gas decreased leading to bubble size expansion.

Figure 6.8b shows the variation in the temperature in the downstream channel (top) and the corresponding variation in the volume of bubbles (bottom), both plotted as a function of the distance from the orifice of the MF device. The stabilization of the bubble size occurred at a distance of ~10 mm from the orifice, which was determined by the saturation of the continuous

o aqueous phase with CO2. Following the reduction in temperature from 25 to 8 C in Region 1 of the downstream channel, the volume of bubbles was reduced by ~30 %, in comparison with their original dimensions. Based on the ideal gas equation and Charles’s Law,40 the contribution from the temperature-dependent volume change of CO2 was estimated to be ~5.1 %. Thus we conclude that the reduction in the volume of bubbles was largely due to the dissolution of CO2 at reduced temperature, rather than a temperature-dependent volume change. A subsequent increase in temperature from 8 to 56 or from 8 to 77 oC in Region 2 led to the 52.5 % or 83.3 % increase in bubble volume, respectively. Next, cooling of the continuous phase to 10oC led to the increased solubility of CO2 in water and the subsequent 34.8 % and 37.7 % reduction in the bubble volume, respectively.

The changes in bubble dimensions occurred within 10 s and did not affect the distribution in bubble sizes. The polydispersity of the bubbles was below 5 %, owing to the uniform dissolution and mass transfer of CO2 between the gas and the liquid phases.

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In the next step, we applied the approach for the temperature-controlled solubility of CO2

‘naked’ bubbles, to the formation of 186 µm CO2 bubbles in 1.2 wt. % dispersion of anionic 2.8

µm-diameter poly(styrene-co-acrylic acid) (PS-co-PAA) particles in a 1M NaOH aqueous solution (pH = 14). We use this value of pH for two reasons. Firstly, at pH=14, the carboxylic groups on the surface of the polymer microbeads were deprotonated, since the value of pKa of

41 the carboxylic groups is 4.25. Secondly, the dissolution rate of CO2 at pH=14 is higher than at lower pH values such as at pH = 6.31 The bubbles were generated at 22 oC and were subjected to a moderate increase in temperature to 28 oC when they traveled through the downstream channel.

Figure 6.9a shows the resulting CO2 bubbles coated with a close-packed shell of polymer

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particles. The particles formed a two-dimensional colloid-crystal layer. The pH reduction induced by the rapid dissolution of CO2 caused the protonation of the carboxylate groups on the surface of the particles in the neighbourhood of the bubble surface and an increase of the wetting angle of water on the polymer surface. The protonation of the carboxylic groups of the polymer particles occurred only close to the surface of bubbles, whereas in the rest of the continuous phase, the particles did not aggregate or precipitate. The deposition of the colloidal shell prevented further dissolution of CO2 from the bubble, due to the reduced interfacial area between the gaseous and liquid phases. Figure 6.9b shows a histogram of the size distribution of armoured bubbles with an average diameter of 60 µm and a narrow size distribution of sizes (CV

= 4.6 %).

In the next series of experiments, we examined the temperature-mediated variation in the volume of armoured bubbles. We varied the temperature of the continuous phase from 1 oC to 77 oC using the same approach as described in section 5.3. Figure 6.10a shows the relative change in the volume of the bubbles dispersed in the 1.2 wt. % dispersion of PS-co-PAA particles in a 1M

NaOH aqueous solution (pH = 14), plotted as a function of temperature. For the same initial dimensions of CO2 bubble of 186 µm, the reduction in the volume of the bubbles was 98.9 % and 81 % at T = 1 oC and T = 77 oC, respectively. Figure 5.10b shows the variation in the diameter of particle-coated bubbles, plotted as a function of the temperature of the liquid phase.

The mean diameter of the armoured bubbles increased from 43 to 115 μm, when the temperature in the downstream channel increased from 1 to 77 oC, respectively.

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The relative change in dimensions of CO2 bubbles was governed by two coupled processes: the temperature-dependent dissolution of CO2 through uncovered segments of the gas-liquid interface and the adsorption of particles to the gas-liquid interface (which reduced the surface area available for CO2 dissolution).

The radius Rb(t) of a bubble and the number of adsorbed particles np(t) can be approximately

31 described by equations 6 and 7, for Rb(t) >> 휶

(6)

(7)

where a is the radius of absorbed particles, A is the uncovered bubble area, α is a constant that accounts for the temperature-dependent gas volume change based on the ideal gas law, qb and qp are the gas and particle fluxes. We expect that qb increases with decreasing temperature, due to the higher solubility of CO2, while qp decreases at reduced temperature, due to the lower

42 diffusion rate of particles at low temperature, following Fick’s diffusion laws. When qp = 0 is the particular case for the dissolution of naked bubbles. The ratio of eqs. 6 and 7 yields:

(8)

where B is a constant at a particular pH, particle concentration, and temperature, which undergoes an increase with decreasing temperature. Therefore, for the initial conditions Rb(0)=R0 and np(0)=0, the radius Req of equilibrium bubble covered with particles radius a satisfies the eq.

(8):

(9)

that is, Req decreases monotonically with increasing B. Therefore, the dimensions of the armoured bubbles decrease as the temperature decreases.

135

The dissolution of CO2 was dramatically slowed down once the bubbles were fully covered with the particles. The dimensions of the armoured bubbles stayed constant due to the formation of a rigid shell composed of jammed particles, which protected the bubbles from coalescence.

6.5 Conclusions

The MF temperature-controlled 'bubble breathing' paves the way for high-throughput studies of dynamic, temperature mediated CO2 interactions with liquid media. The ability to generate bubbles with narrow size distributions solves the problem of bubble disparity in size, which constitutes a problem in conventional studies of CO2.

The ability to monitor the difference in temperature-controlled dissolution of CO2 in deionized water and ocean water has important implications for environmental sciences. We also showed the 'breathing bubble' effect in DEPG, a physically absorbing solvent. The proposed MF method can be used as an efficient exploratory tool. A high-throughput variation in the amount of CO2 and a particular reagent(s) supplied to the system at varying temperatures, followed by the evaluation of the amount of reacted or physically sequestered

CO2 can be used for the development of new formulations and optimization of conditions for efficient CO2 sequestration. Ocean water contains a wide range of organic compounds

(commonly refered to as volatile organic compounds), which may act as surfactants and reduce the surface tension at the gas-liquid interface, therby affecting the transfer of gas to the liquid. Therefore, the MF approach described in the present chapter can be useful for monitoring the effect of surfactant on the dissolution of gases in various solvents.

For reversible reactions, e.g., “solvent polarity switches”, the MF method enables the determination of temperatures at which a reverse reaction leading to CO2 release becomes dominant. For physical sequestration of CO2, this method allows for optimization of reaction

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conditions such as temperature for sequestration by solvents such dimethyl ethers of poly(ethylene glycol).

The MF approach can be utilized for studies of other highly reactive gases, e.g., H 2S, or SO2, with temperature-dependent solubility in liquid media. These gases have important environmental impact, owing to their high anthropogenic generation and their contribution to the greenhouse effect.43 Reactive gases are also used in heterogeneous gas–liquid chemical reactions.44 Monitoring temperature-controlled variation in bubble dimensions paves the way for the fundamental studies of these reactions, as well as their kinetics and thermodynamics.

Further advancements in MF studies of CO2 will include the use of automated processes to monitor and determine bubble dimensions under a broad range of conditions. 45 This method would enable extremely high-throughput studies of kinetics and thermodynamics of gas/liquid reactions. The modification of the MF device with extended microchannel length would allow bubbles to reach their equilibrium size. A deeper understanding of the CO 2 mass transfer at the gas-liquid interface is highly desirable to further ascertain fundamental knowledge of these reactions.

Furthermore, it was demonstrated that temperature-controlled dissolution of CO2 can be used for the preparation of highly monodispersed particle-encapsulated bubbles. By generating CO2 bubbles at room temperature and changing the temperature of the continuous phase in the downstream channel, we achieved control over CO2 dissolution, thereby producing monodisperse bubbles with a controllable final size. The size of armoured bubbles can be further decreased by using a modified droplet generator and using a higher flow rate of the continuous phase.21 The MF approach can be extended to droplets carrying a mixture of several components, one of which exhibits the temperature-dependent solubility in the continuous phase.

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The proposed approach is useful for the generation of a large range of functional bubbles or droplets coated with nano and micro particles for applications such as magnetic resonance imaging (MRI) and drug delivery.

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(20) K. Hettiarachchi, E. Talu, M. L. Longo, P. A. Dayton and A. P. Lee, Lab Chip 2007, 7, 463.

(21) J. I. Park, Z. H. Nie, A. Kumachev and E. Kumacheva, Soft Matter 2010, 6, 630.

(22) U. T. Gonzenbach, A. R. Studart, E. Tervoort and L. J. Gauckler, Angew. Chem.-Int. Edit., 2006, 45, 3526.

(23) B. P. Binks and R. Murakami, Nat. Mater. 2006, 5, 865.

(24) Z. P. Du, M. P. Bilbao-Montoya, B. P. Binks, E. Dickinson, R. Ettelaie and B. S. Murray, Langmuir 2003, 19, 3106.

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Chapter 7 Development and Applications of Microfluidic Reactors

with Multiple Reconfigurable Analytical Probes

7.1 Introduction

Chemical synthesis in microfluidic (MF) reactors is highly efficient due to the excellent control over reaction conditions, low reagent consumption, precise timing between subsequent reaction steps and the ability to isolate reactive intermediates from ambient conditions.1-5 These features make MFs an ideal tool for optimisation of formulations and reaction conditions.

Future advancements in the applications of MF chemical synthesis are largely determined by the ability to carry out in situ characterisation of the chemical and physical changes in reagents and products, as well as monitoring reaction conditions, such as changes in temperature, pH, ionic strength, and pressure.2 Advances in the integration of chemical characterisation tools with MF systems include the development of various kinds of in situ analytical tools such as NMR,6,7 fluorescence,8 infrared (IR),9,10 Raman spectroscopies,11 surface enhanced Raman spectroscopy (SERS),12 and pH probes.13,14 Yet, generally chemical characterisation in MF systems is conducted by interfacing a MF reactor with a particular single probe, in order to characterise a well-defined reaction variable, whereas the integration of several in situ probes can provide complementary information about several reaction variables. For instance, reactions including consumption or production of mineral acids such as HCl, cannot be solely monitored by IR spectroscopy; however, complementary in situ pH measurements can enable a direct measurement of the change in the concentration of the 142

acid. In addition, the use of complementary probes provides the ability to explore a large number of reaction variables in parallel, which is useful for the optimisation of chemical formulations. To the best of our knowledge, the only report of the integration of multiple characterisation techniques in a MF bioreactor system was employed to monitor biological processes.15 This MF system was utilized for continuous cell culture, however, HPLC-based measurements of the culture media and temperature measurements were conducted off-chip, that is, not in situ; and the pH measurements required additional off-chip verification using a pH electrode, due to the drift associated with photobleaching. In principle, fluorescence- based molecular probes can be devised for monitoring several reaction variables in parallel; however this strategy is limited by the overlap of spectral features of multiple probes.

Importantly, the utilisation of exchangeable integrated probes would offer the capability of using the same MF reactor for studies of many different chemical reactions.

In this chapter, we report the development and applications of a versatile MF platform with multiple reconfigurable in situ analytical probes that can be used for (i) measurements of acidity and basicity of the reaction system, (ii) the quantitative characterisation of molecular vibrational signatures of reactants or products, and (iii) the real-time monitoring of temperature. The MF reactor was fabricated in polycarbonate (glass transition temperature of

145 oC) by thermoembossing,16 The stamp was produced by a photolithographic method. The analytical tools included an Attenuated Total Reflection Fourier Transform Infrared (ATR-

FTIR) spectroscopy probe, a miniaturised pH electrode, and a thermocouple element, with the pH and temperature probes being interchangeably embedded into the MF reactor. The integration of these probes was sufficiently robust to be used at 0 ≤ pH ≤ 14 and a temperature range from 0 to 100 oC. (The upper temperature can be increased, as it is limited

143

by the glass transition temperature of the thermoplastic material of the reactor). The temperature and pH probes were reconfigurable: a particular MF reactor could be interfaced with interchangeable temperature and pH probes, without the need for re-design of the MF reactor. In addition, if required, the reactor could be integrated with two pH or two temperature probes. Furthermore, we show the ability to use the temperature probe as a feedback system to control the temperature in the MF reactor.

The rationale in selecting the ATR-FTIR, the pH, and the temperature probes was as follows: the utilisation of ATR-FTIR spectroscopy allows full-spectrum, qualitative and quantitative characterisation of chemical and biological species, without the need for sample preparation or the addition of probe molecules. Since the effective path length of ATR-FTIR measurements is below 2 µm,17 data acquisition can be readily achieved in highly absorbing aqueous environments that would completely attenuate the signal in transmission measurements. Recently, in situ ATR-FTIR has been used for the spectroscopic characterisation of soluble analytes and molecules adsorbed to the surface of the ATR crystal.9 Measurements of site-specific pH values are beneficial for MF reactions that undergo a change in pH,18 or are dependent on the pH of the medium.19 Measurements of the temperature of the reaction system enable the studies of temperature-dependent reaction mechanisms. Monitoring the generation or consumption of heat can also provide useful information on reaction thermodynamics, such as G or equilibrium constants.

The multiprobe MF reactor reported in the present work had the following useful characteristics: (i) the ability to characterise a site-specific (and hence time-specific) state of the system, (ii) the ability to reversibly remove or exchange the pH probe and the temperature probes and (iii) integrated feedback for a heat-controlled circuit for temperature

144

control; and (iv) the fabrication of the MF reactor in a robust, optically transparent thermoplastic material enabled an additional mode of system characterisation by optical microscopy.

7.2 Experimental Design

7.2.1 Integration of Probes with the MF Reactor

Figure 7.1 a and b shows a schematic of the MF reactor interfaced with the ATR-FTIR, pH and temperature probes. The probes were located along the MF reactor at points P1, P2 and

P3, allowing for site-specific and hence time-specific measurements at different stages of the reaction (Figure 7.1a). The liquid reagents were supplied to the reactor via two inlets and were subsequently mixed in a serpentine channel. A probe located at point P1 between the mixing and reaction compartments provided information about the state of the system at the beginning of the reaction. Points P2 and P3 were located between the end of the reaction compartment and the outlet, allowing characterisation of the system after the reaction was completed. As noted in Figure 7.1a, the positions of the pH, temperature, and IR probes were arbitrary and were determined by the requirements of a particular reaction system. Moreover, it was possible to introduce two temperature or two pH probes to characterise the initial and final states of the reaction. Figure 7.1c shows an optical microscopy image of the fragment of the microchannel passing along the surface of the circular ATR crystal. The MF reactor was immobilised over-top of the ATR-FTIR accessory. A photograph of the packaged MF reactor with integrated probes and fluidic connections is shown Figure 7.1d.

We examined the following characteristics of the integrated probes: (i) the response time of each individual probe to a change in reaction conditions, (ii) the resistance to signal drift of each probe with increasing flow rate of the continuous phase, (iii) the reproducibility and

145

absence in hysteresis when the concentrations of analytes are changed by varying the flow rates of the liquids.

7.2.2 Validation of Measurements of pH

As shown in Figure 7.2a, the pH probe was introduced in the MF reactor at position P2. In

146

order to ensure that the response of the pH probe is not affected by the shear stress exerted by the flow of the liquids or by the variation in pressure accompanying changes in flow rates of the liquids, a solution of HCl was supplied to the MF reactor at varying flow rates. Figure

7.2a shows that the value of pH = 2.3 was measured at t = 0 when the solution of HCl was supplied in the MF reactor (Pump-1 on and Pump-2 off). At t = 10 s the supply of the liquid was switched from Pump-1 to Pump-2, thereby injecting in the reactor a solution of NaOH.

An increase in basicity to pH = 11.0 was measured 15 s after Pump-1 was turned on. The lag time corresponded to the time it took for the base solution to travel through the microchannel and reach the probe at point P2. At t = 90 s, Pump-1 to Pump-2 were switched, again, so that the flow of the NaOH solution was stopped and the flow of the HCl solution was resumed.

With a 15 s delay, a sharp decrease in acidity to pH = 2.5 occurred. The time for the pH to change from 2.3 to 11.5 and from 11.5 to 2.3 was ~60 s. Two cycles of the change in pH are shown in Figure 7.2a and were representative of 5 cycles. Figure 7.2b shows that in the range of flow rates from 0 to 20 mL/h the value of pH of the solution did not change. The response time and reversibility in pH measurements were examined by supplying in the reactor in an alternating manner an acidic and a basic solution. We used two syringe pumps (Pump-1 and

Pump-2, respectively) to introduce in the MF reactor an aqueous solution of HCl (pH = 2.3) or an aqueous solution of NaOH (pH = 11.5). The flow rate of each solution was 2 mL/h.

Figure 7.2b shows the results of pH measurements. In the range of volumetric flow rates of the liquid of up to 20 mL/h we did not observe any change in the value of pH.

147

To verify that the lag in response time was dominated by the inherent response time of the pH probe and not the mixing time of the acid-base solutions at the interface between the two liquid streams in the MF reactor, we conducted a control experiment by transferring a pH probe from a bulk acidic solution (pH = 2.3) to a bulk basic solution (pH = 11.5). The time required for the pH to increase was ~40 s, in accordance to the specifications provided by the probe manufacturer and sufficiently close to that measured in the MF experiments.

7.2.3 Validation of FT-IR characterization

9 As shown in Figure 7.1a, an ATR probe was interfaced with the MF reactor at point P2. We supplied to the MF reactor two liquids: (i) deionised water and (ii) a 50 mM of

148

tris(hydroxymethyl)aminomethane (Tris) buffer solution (titrated at 25 oC with 1.0 M HCl to achieve pH = 7.15). The liquids were introduced in an alternating manner, similar to that described in the preceding section. The response time of the probe was characterised by monitoring changes in the infrared absorbance of Tris. We monitored the change in the intensity of the band at 1060 cm-1, which we assigned to the Tris C-N stretching band (Figure

7.3c and d).20

Figure 7.3a shows the response time and the reaction reversibility, both were determined by using ATR-FTIR measurements, when either the Tris buffer solution or deionized water were supplied to the MF reactor. At t = 10 s, the supply of deionized water was turned off and the

Tris buffer supply was turned on. Within the time 20 ≤ t ≤ 40 s absorbance of the Tris C-N band increased; between 40 < t ≤ 85 s, the absorbance signal stabilized. At t = 75 s, the supply of Tris solution was replaced with deionized water and a subsequent decrease in the

C-N absorbance was observed. This process was repeated for 5 cycles. The response time to the change in the intensity of the absorbance signal was consistently 30 s in length.

Next, we ensured that the absorbance measurements were not affected by variation in the flow rate of Tris solution. We introduced a 50 mM Tris buffer solution (pH = 7.15) into the

MF reactor at flow rates in the range of 0 < Q < 20 mL/h. Figure 6.3b shows absorbance data for a band at 1060 cm-1, collected using an ATR-FTIR probe. No change in absorbance occurred in the designated flow rate range.

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Figure 7.3 (a) Variation in absorbance measured for the 1060 cm-1 band following injection of water and 50 mM Tris solution. Spectra were acquired based on a single measurement comprising 4 scans at 10 kHz and 8 cm-1. The time between the dashed lines indicates the time required for the absorbance at 1060 cm-1 to change (~ 15 s). Arrows indicate the time of stopping of the flow of either the water, or the Tris solution and the simultaneous introduction of the other liquid. Each solution was supplied at the flow rate Q = 3 mL/h (b) Variation in absorbance (1060 cm-1) vs. flow rate of the 50 mM Tris solution. Error bars were obtained by determining the standard deviations in absorbance, based on five independent measurements for each flow rate of the liquids. Spectra were collected using 8 scans at 10 kHz and 8 cm-1 spectral resolution.

7.2.4 Validation of Temperature Measurements

As shown in Figure 7.1a, a thermocouple connected to a digital thermometer was interfaced with the MF reactor at point P1. The temperature in the reactor was varied between 25 and 50 oC using a heating pad placed below the bottom of the reactor. The device was cooled by placing it an ice bath. In this system, the response of the system was not limited by the probe response, but instead, by the time required to heat or cool the entire MF system, which includes the heating pad, the thin sealing layer of the MF device, and the liquid flowing through the microchannel. Deionized water was supplied to the reactor at a flow rate of 4 mL/h and a temperature of 20 oC. Figure 7.4a shows the variation in temperature of the water

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flowing in the microchannel with respect to time. The change in temperature was controlled by setting a specific temperature with the temperature controller.

Figure 7.4a shows two heating and cooling cycles. At time 20 s, the heating pad was activated and the temperature was set to 50 oC. Within ~500 s the liquid was heated from 25 to 50 oC. Upon reaching the target temperature of 50 oC, the temperature was maintained constant without any detectable variation in temperature. At this time, the temperature of the heating pad was changed to 25 oC. The temperature controller attenuated current supplied to the heating pad and the flow of heat to the ice bath placed below the device led to the reduction in the temperature of the liquid stream. The controller supplied sufficient current to the heating pad, in order to maintain the 25 oC set point. We achieved a minimum time to heat the flowing water in the microchannel from 25 to 50 oC in ~120 s. This was accomplished by adjustment of the PID (proportional–integral–derivative) settings of the temperature controller. In this case, the stabilisation of temperature was preceded by a 60 s time interval within which the temperature oscillated due to an overshoot of 5 oC (not shown in Figure 7.4a).

To determine the time required for a liquid at an initial temperature of 20 oC to reach a target

o temperature of 50 C prior to reaching point P1, we conducted experiments at varying flow rates of water. The temperature of the heating pad was set and maintained at 50 oC. Figure

o 7.4b shows that at flow rates below 3.5 mL/h, the temperature of the liquid at P1 was 49 C.

We assumed that a 1 oC difference between the set temperature and the measured temperature of the water was the result of the temperature gradient across the bottom layer of the reactor. At flow rates exceeding 3.5 mL/h, the liquid did not have sufficient time to reach

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the equilibrium temperature prior to reaching position P1 (Figure 7.4b, circles) and the measured temperature was consistently reducing with increasing flow rate of the liquid.

We supported experimental results with simulations of heat transfer from the heating pad to the liquid flowing through the MF reactor. We modelled the expected temperature of the liquid traveling at flow rates in the range from 0 to 10 mL/h through the entire reactor at the temperature of the heating pad of 50 oC. Figure 7.4b (dotted line) shows the results of the simulations of the temperature of the liquid at P1 were in agreement with experimental results. Furthermore, at flow rates of up to 10 mL/h, simulations predicted that the liquid reaches the target temperature of 50 oC before entering the reaction compartment which was located 3.5 cm downstream from the inlet. Figure 7.4c shows an exemplary simulation result for water flowing at r.t. into the MF device at a flow rate of 0.9 mL/h. At the flow rate of 0.9 mL/h, the temperature of the flowing liquid reaches thermal equilibrium at ~ 2 mm beyond the orifice, which is a significant distance from the first probe point.

To ensure that the liquid attained the target temperature at point P1, irrespective of its flow rate, we implemented a feedback temperature control of the liquid based on the real-time temperature measurements at point P1. The feedback approach enabled an automatic adaptation of the thermal output of the heating pad, in order to compensate for the short residence time of the liquid moving at high flow rates through the MF reactor. The feedback system allowed us to reach the target temperature and maintain it at a precisely controlled value at flow rates from 0 to 10 mL/h (Figure 7.4b), that is, well above 3.5 mL/h, which limited the flow rate for the system without feedback temperature control. Furthermore, we maintained control of the temperature at 50 oC at flow rates of up to 50 mL/h (not shown in

Figure 7.4b).

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7.3 Applications of the MF Reactor Integrated with Multiple Probes

7.3.1 Titration Reactions

Titration is an analytical method for quantitative chemical analysis. Titration reactions are used in (i) acid/base neutralization,21 (ii) redox reactions,22 and (iii) chelation reactions.23

Generally, titration involves the determination of the concentration of an analyte (the titrand), by adding a titrant with a known concentration. As an example, an acid/base titration allows the determination of an unknown concentration of one of the starting materials by administering the titrant until the equivalence point is reached. In the case of acid-base titrations, the equivalence point is reached when the acid and base solutions become mutually neutralized after mixing.

7.3.2 Titration in a Microfluidic Format

When the acid and base solutions are mixed, the initial concentrations of the acid and the base reduce to [A]d and [B]d, respectively, as in

[A]d = [A]iQA/QT (1a)

[B]d = [B]iQB/QT (1b)

The extent of the neutralization in a titration experiment depends on the values of [A] d and

[B]d. Furthermore, a titration can reveal several equivalence points. For example, the titration of an acid containing a specific number of acidic moieties per molecule (a multiprotic acid), with NaOH (a monoprotic base) will reveal the same number of equivalence points, each corresponding to the full neutralization of one acid moiety. This is discussed in section 7.3.5

Titration of a diprotic acid. For a monoprotic acid titrated with a monoprotic base, the

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equivalence point is reached when [A]d = [B]d. Based on eq. 1, we obtain:

[A]i  QA,e = [B]i  QB,e (2)

-1 - where [A]i and [B]i are given in mol L , and QA,e and QB,e are the volumetric flow rates (L s

1) of the acid and the base solutions, respectively, required to reach the equivalence point.

The ratio of flow rates of the acid and base solutions at which the system achieves molar equivalence is determined by the initial concentrations of the corresponding solutions as in:

QB,e/QA,e = [A]i/[B]i (3)

For [A]i/[B]i=1, the molar equivalence is achieved for QB,e/QA,e=1. By independently modulating the flow rates of each solution, the equivalence flow rates will be reached when this work, the total flow rate of the acid and base solutions was maintained constant (QT =

QA + QB = constant). The equivalence flow rate of the acid solution is QA,e = QT – QB,e and eq. 3 can be solved for QB,e as:

QT [A]i /[B]i Q  (4) B,e (1[A] /[B] ) i i

In the titration experiments described here, the value of pH was measured for different ratios

QB/QT. Therefore, QB/QT is the experimental variable and pH is the measured output. To demonstrate a proof of concept, up to 40 evenly spaced pH values were acquired in the range of 0 ≤ QB/QT ≤1. Typically, a larger number of points were acquired for the regions of the titration curve which represented the region near the equivalence point and had a large slope.

Using the integrated temperature probe (inserted at point P2) we monitored the temperature after the acid and base solutions underwent mixing/neutralization. Additional temperature

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control could be accomplished by using a planar heating pad located below the MF device, with a feedback control from the temperature probe. This could be optionally implemented when the neutralization experiments were conducted at temperatures different than r.t. conditions.

7.3.3 Titration of a Strong Acid with a Strong Base

The chemical reaction between a strong acid and a strong base is given by:

+ - 14 H + OH ⇄ H2O, K1 = 10 (5)

where K1 is the equilibrium constant for this reaction (K1 = 1/Kw where Kw is the self-

24 ionization constant for water). The large value of K1 indicates that the reaction goes to completion, and the acid reacts in a stoichiometric equivalent of the base.

We used the solutions of hydrochloric acid (HCl) and sodium hydroxide (NaOH) as the strong acid and strong base, respectively. In three separate experiments we introduced solutions into the MF reactor at the initial concentrations of the HCl solutions, [HCl]i =

-1 0.025, 0.035 and 0.055 mol L , and the initial concentration of NaOH solution, [NaOH]i =

0.05 mol L-1.

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Firstly, using eqs. 4, 6 and 7 we determined the theoretical value of pH in three regions of the titration curve, namely, before the equivalence point (QNaOH/QT < QNaOH,e/QT), at the equivalence point (QNaOH/QT = QNaOH,e/QT) and after the equivalence point (QNaOH/QT >

QNaOH,e/QT). In the region before the equivalence point, the excess HCl is given by:

+ [H ]f = [HCl]d – [NaOH]d (6a)

Q Q [H  ]  [HCl]  HCl  [NaOH]  NaOH (6b) f i Q i Q T T

where the multiplication of the initial reagent concentrations, [HCl] i and [NaOH]i, with their

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Q Q respective volume fractions, HCl and NaOH , yields the concentrations of the reagents QT Q T upon dilution (eq. 1). After the equivalence point, the excess of NaOH is given by:

- [OH ]f = [NaOH]d – [HCl]d (7a)

 Q NaOH Q HCl [OH ]  [NaOH]   [HCl]  (7b) f i Q i Q T T

+ + - The value of pH can be determined from eqs. 6 and 7 as -log[H ] (for eq. 7b [H ]=Kw/[OH ]).

At the equivalence point, pH = 7, since a strong acid and strong base undergo complete neutralization with each other to produce water.

Table 1 summarizes the initial concentrations and pH values of the reagent solutions, along with the calculated and experimentally measured ratios of QNaOH,e/QT. The experimental and calculated titration curves of QNaOH/QT in the range of 0≤QNaOH/QT ≤ 1, at different concentrations of [HCl]i is shown in Figure 7.5.

In Figure 7.5, all the plots begin at low QKOH/QT, resulting in acidic conditions. As the ratio

QKOH/QT approached the value required for the equivalence point, a sharp increase in the pH value was measured. The pH curves calculated using eqs. 6 and 7 are plotted on the same graph as the dashed lines. The theoretical and experimental data were in excellent agreement.

NaOHi HCli QNaOH,e/QT QNaOH,e/QT [NaOH]i pH [HCl]i pH (calculated) (measured) (mol/dm3) (mol/dm3) 0.050 12.7 0.025 1.60 0.32 0.32 0.050 12.7 0.035 1.50 0.41 0.41 0.050 12.7 0.055 1.30 0.51 0.52

Table 7.1 Summary of the Titration of a strong acid (HCl) and a strong base (NaOH)

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7.3.4 Titration of Weak Acid with Strong Base

We used KOH and CH3COOH as a strong base and a weak acid, respectively. The reaction occurs according to:

- - 9 CH3COOH + OH  CH3COO + H2O K2 = 1.61 x 10 (8)

where K2 is the equilibrium constant for the reaction. This reaction is a reverse reaction of the association of a conjugate base with a proton (with equilibrium constant Kb). Therefore

24 K2 = 1/Kb = Ka/Kw, where Kb = Kw/Ka. Since CH3COOH is a weak acid, it does not fully dissociate in water and the proton concentration is given by:

- + CH3COOH ⇄ CH3COO + H pKa = 4.8 (9)

We introduced the acid and base solutions into the MF reactor at initial concentrations

[CH3COOH]i = 1M and [KOH]i = 1M. The calculated pH of the initial acid and base solutions were 2.4 and 14, respectively.22 We analyzed the titration curve in three regimes, namely, QKOH/QT < QKOH,e/QT (before the equivalence point), QKOH/QT = QKOH,e/QT (at the equivalence point) and QKOH/QT > QKOH,e/QT (after the equivalence point). The high value of the equilibrium constant in eq. 8 indicates that before the equivalence point the reaction goes

- to near completion. Thus, more acetate ions (CH3COO ) molecules will be created (and an equivalent amount of CH3COOH consumed) in molar equivalence with the amount of KOH

- added to the system. Since the mixture of CH3COOH and CH3COO forms a buffer, the pH value can be calculated using the Henderson-Hasselbalch equation as:24

 [CH COO-]  pH  pK  log 3 f  (10) a [CH COOH]   3 f 

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- where [CH3COOH]f and [CH3COO ]f are the final concentrations of CH3COOH and its conjugate base after neutralization with KOH, respectively. Since before equivalence KOH is fully consumed, using eq. 3, we obtain:

[CH3COOH]f = [CH3COOH]d – [KOH]d (11)

Equation 10 can therefore be re-organized as:

 QKOH   [KOH]i    Q  pH pK  log T (12) a  Q  CH3COOH QKOH [CH3COOH]i  [KOH]i    QT QT 

In the experiments, we used [KOH]i = 1M and [CH3COOH]i = 1M. It follows from eq. 10

- that pH = pKa when [CH3COOH]f =[CH3COO ]f. Since at this point, the buffering capacity of the solution comprised of acetic acid and its conjugate base is the strongest, the slope of the plot of pH vs. QKOH/QT will be at a minimum, and this point can be detected as an inflection point on the titration curve. For QT = 2mL h-1, this point should occur for

-1 -1 QKOH=2/3 mL h and QCH3COOH,e = 4/3 mL h . Therefore, at a flow rate ratio of QKOH/QT =

-1 1/3 mL h we expect pH = 4.8. The predicted value of pH at the equivalence point is pHe =

9.3.24

In the MF titration experiments, the acid and base solutions passed through the mixing region in the MF device in ~10 s and subsequently entered the pH probe location (P1). The titration curve was acquired by measuring pH values at varying ratio QKOH/QT (Figure 7.7). Figure 7.7 shows the following features: (i) an initial fast pH increase for Q KOH/QT < ½QKOH,e/QT, (ii) an inflection point at QKOH/QT = ½QKOH,e/QT and pH = pKa, (iii) the equivalence point at

QKOH/QT = QKOH,e/QT, and (iv) the asymptotic approach to the value of pH of the pure KOH 160

for QKOH/QT > QKOH,e/QT. Inspection of Figure 7.7 showed an inflection point at QKOH/QT =

0.3, in agreement with the predicted ratio of QKOH/QT = 1/3.

Figure 7.6 Microfluidic titration of a weak acid with a strong base. (a) Titration curve for [CH3COOH]=1M, [KOH] =1M. Data acquisition was conducted 2-3 min after changing the flow rates of the liquids. The pKa is measured from the first inflection point (where the change in pH is minimum) and the pHe is measured from the second inflection point (where the change in pH is maximum).

The pH value measured under these flow conditions was approximately 4.4, compared to 4.8, the value of pKa for CH3COOH. Since [KOH]I = [CH3COOH]I =1 M, the equivalence flow rate ratio was predicted to be QKOH,e/QT = 0.50 (eq. 3). Based on the data in Figure 7.7, we determined pHe = 9.3, estimated from the location of the inflection point on the pH curve

(QKOH,e/QT = 0.54).

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7.3.5 Titration of a Diprotic Acid

The titration of a diprotic acid results in two equivalence points, each corresponding to the neutralization of one acid moiety. The first equivalence point (the neutralization of the lower

pKa acid moiety at pKa1) is achieved when the concentration of the acid, [A]d, and base,

[B]d, are present in molar equivalence, analogous to the neutralization of a monoprotic acid

(eq. 3). Thus, the flow rates of the acid and base solutions leading to neutralization of the first acid moiety are determined by eq. 4. After the neutralization of the first acid moiety, the concentration of the base has to be increased to reach the second equivalence point, at which the second acid moiety is neutralized. Reaching this point requires approximately twice as many base molecules per diprotic acid than were required to neutralize the first acid moiety.

Therefore, full neutralization of the higher pKa acid moiety (pKa2) is achieved for [B]d 

2[A]d.

We used KOH and malonic acid, CH2(COOH)2, a diprotic acid with pKa1 = 2.85 and pKa2 =

5.70.24 We introduced the acid and base solutions into the reactor at initial molar concentrations [CH2(COOH)2]i = 0.09M and [KOH]i = 1.00 M. The initial pH value of the

0.09M CH2(COOH)2 solution was measured to be 1.95.

The experimental results are shown in Figure 6.8a in the range 0 < QKOH/QT < 0.250. In this flow rate range, the value of pH was rapidly changing. As discussed in the previous section, the Henderson-Hasselbalch equation implies that the slope of the dependence of pH vs.

24 QKOH/QT is at a minimum when pH = pKa. For multiprotic acids, we expect to see multiple slope minima and several equivalence points on the titration curve, as shown in Figure 7.8a.

While the values of pKa and pHe could be acquired by eye, we used a first derivative plot of the pH vs. QKOH/QT to accurately determine the flow rates of the liquids, which corresponded 162

to these pH values (Figure 7.8b). The slopes of the lines connecting each data point were determined using standard spreadsheet software (Microsoft Excel 2010) and plotting them against QKOH/QT. From Figure 7.8b, we identified the local minima at QKOH,pKa1/QT = 0.069 and QKOH,pKa2/QT = 0.131. These points corresponded to the measured pH values of 2.80 and

5.65, respectively, which were close to the reported pKa1 = 2.85 and pKa2 = 5.70 for

CH2(COOH)2. This advanced approach can be used for in-depth studies of multiprotic acids or for mixtures of several acids, as well as base mixtures.

Figure 7.7 Microfluidic titration of a strong base with a multiprotoic acid. (a) Titration curve for [CH2(COOH)2] = 0.09M, [KOH] = 1.00M. The data were acquired based on the two independent experiments. Acquisition was conducted 2-3 min after changing the flow rates of the liquids. The values of QKOH/QT yielding pH = pKa1 and pH = pKa2 were determined by eye at the inflection points in on the titration curve, where the slope was lowest. The values of QKOH/QT corresponding to pH = pHe1 and pH = pHe2 were determined by eye at the inflection points where the slope was largest. (b) The exact QKOH/QT values yielding pH = pKa1 and pH = pKa2 were determined by the local minima in the first derivative plot of (a). The exact QKOH/QT values leading pH = pHe1 and pH = pHe2 are determined by local maxima in the first derivative plot of (a).

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7.3.6 Titration of a Strong Acid with a Strong Base with Simultaneous On-Chip Temperature Measurements

To demonstrate the simultaneous data acquisition and characterisation of two reaction variables by integrating the temperature and the pH probes, we performed an on-chip neutralisation reaction at varying concentrations of a strong acid and a strong base solution.

Similar to the previous section, we introduced in a MF reactor two aqueous streams: a solution of strong acid, HCl at pH = 0.3 and a solution of strong base, NaOH at pH = 13.4.

We maintained a constant total volumetric flow rate of the two liquids at 4 mL/h and changed the relative flow rate ratios of the acid and base solutions. The solutions were introduced in the reactor at 23 oC. The temperature and pH of the liquid stream were measured at points P1 and P2, respectively. Figure 7.6 shows that when the ratio of the molar concentrations of the HCl-to-NaOH solutions changed from 1:0 to 1:1, the increase in temperature reached a maximum value of 1.4oC. Subsequently, when the molar concentration ratio was reduced to [HCl]:[NaOH] = 0:1, the temperature of the solution reverted back to the original temperature (that is, the change in temperature compared to the initial temperature was reduced to 0). The variation in pH exhibited characteristic acid/base titration behaviour with an equivalence point at pH 7.24 Comparison of the variation in pH and temperature of the solution showed excellent correlation: the maximum increase in temperature occurred when the concentrations of HCl and NaOH solutions corresponded to the complete neutralisation of the acid. Since neutralisation reactions involving a strong monoprotic acid and a strong base generate heat, the highest temperature of the liquid was achieved at equal concentrations of the acid and the base.21

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7.4 Temperature-Dependent Change in the pH of Tris Buffer

To demonstrate a relationship between pH and temperature, we examined the variation in the pH values of a solution of Tris buffer as a function of temperature of the liquid medium. Tris is a small, bio-compatible molecule ((HOCH2)3CNH2)), which is used to maintain pH in the physiological range. The Tris molecule has pK values of 7.82 and 8.08 at 37 °C and 25 °C, respectively.25-28 The buffering capability of Tris results from the lone electron pair on the amine group, which abstracts a proton to form a quaternary amine. The regeneration of the amine by the loss of a proton in basic pH conditions allows Tris to maintain a constant pH value following the exothermic reaction:29

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+ - -1 Tris + H2O  TrisH + OH ΔH = -11 kJ mol (13)

In accordance with Le Chatelier's principle, upon heating, the equilibrium shifts towards the reactants,26 which results in a reduction in pH.

We prepared a 50 mM Tris buffer solution and titrated it at 25 oC with 1.0 M HCl to pH values of either 7.90, or 7.15. We used the temperature feedback control system (described in

Section 7.2.4) by inserting a temperature probe at point P1 (Figure 7.1a). Once the liquid achieved the desired temperature, the pH value was recorded at pH meter which was positioned at P2 (Figure 7.1a). Figure 7.9a shows that in the range of temperatures from 22 to

50 oC, the pH value of both buffer solutions linearly decreased with temperature with a slope of -0.03 pH units/oC, which was in agreement with the specifications reported by the supplier of the buffer30 and control experiments performed in bulk solutions using a standard

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laboratory pH probe.

7.5 pH-Dependent Variation in the Intensity of IR Signals

To demonstrate the simultaneous utilisation of ATR-FTIR and pH probes (inserted at points

P2 and P3, respectively, Figure 7.1a), we monitored pH-dependent changes in the intensity of the vibrational bands of Tris. We mixed a 200 mmol/L solution of Tris and 1 mol/L solution of HCl at the ratio of flow rates of Tris-to-HCl solutions varying from 4:0 to 4:4.9. The

o temperature at point P1 (Figure 7.1a) was maintained at 25 C. The range of measured pH values was from 5.1 to 11.2. As pH progressively decreased, the protonation of the amine

+ -1 group on Tris resulted in growth of the δasNH3 absorption band at 1535 cm (asymmetrical

31,32 -1 bending mode) and the reduction of the vC-N + vN-H2 combination band at 1410 cm

(stretch).31,32 Figure 7.10b and c shows pH-mediated changes in vibrational spectra within the spectral windows 1580-1340 cm-1.

7.6 Reaction of Carbon Dioxide with Water in the Presence of Tris

To exemplify the multi-modal MF platform, we utilised the pH and the ATR-FTIR probes

(introduced at points P2 and P3, respectively) to monitor the kinetics of the conversion of carbon dioxide (CO2) to bicarbonate in the presence of Tris, which proceeds according to the following reaction:

- + CO2 + H2O + Tris  HCO3 + TrisH (14)

As a result of this process, Tris is an efficient blood buffer used for treatment of acidaemia,

25-28,33 which can occur due to CO2 retention or metabolic acid accumulation. In contrast to bicarbonate, a traditionally blood buffer, Tris does not produce CO2, which may exacerbate

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acidaemia.28,33

We prepared a 15 mmol/L Tris buffer solution, titrated it to pH = 9.2 with a 1M solution of

HCl and introduced it into the MF reactor as Reagent 1 (Figure 7.1a). The same Tris buffer solution was saturated with CO2 and introduced into the reactor as Reagent 2. ATR-FTIR measurements of Reagent 2 showed the presence of dissolved CO2(aq) after saturation. After mixing Reagents 1 and 2, the reaction of CO2 (aq) with water was followed by monitoring the intensity of the band at 2342 cm-1 which corresponded to asymmetric O-C-O stretching.

Based on reaction (14), we expected a reduction in the absorbance of the 2342 cm-1 peak. To monitor the kinetics of reaction (14), we modulated the time (t) before the mixture reached point P3 (ATR-FTIR probe) as:

t = D × A/(Q1+Q2) (15) where D and A are the length and cross-sectional area, respectively, of the microchannel in

-1 which the reaction takes place, and Q1 and Q2 (mL s ) are the volumetric flow rates of streams 1 and 2. In the experiments we maintained Q1 = Q2. Figure 6.10 a and b show the variation in CO2 (aq) absorbance and pH with reaction time. To eliminate the effect of temperature on CO2 solubility and on buffer pH, we verified that the temperature of the system remained constant by using a temperature probe interfaced with the MF reactor at

-1 point P1 (Figure 6.1a). Figure 6.10a shows that the absorbance peak at 2342 cm from dissolved CO2 (aq) decreased exponentially

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-1 Figure 7.10 (a) Variation in absorbance of the 2342 cm absorption band of CO2 (aq) plotted as a function of time after mixing Reagents 1 and 2. Each absorbance data point was taken from an individual spectrum comprising 48 scans at 10 kHz and spectral resolution 4 cm -1. Background measurements using deionized water were conducted between experiments. (b) Variation in pH of Tris buffer after introducing CO2 in the liquid phase, plotted as a function + of time after mixing Reagents 1 and 2. (c) Temporal variation in [CO2] and [H ] plotted using the data shown in (a) and (b). (d) FTIR spectrum of 50 mM Tris buffer solution in water (pH = 9.2) flowing through the MF reactor at the volumetric flow rate Q=1 mL/hr. The band highlighted at 1060 cm-1 (indicated with an arrow) corresponded to the C-N stretching vibrational band. (e) FTIR spectrum of CO2 (aq) in Tris buffer (pH = 6.4). The peak at 2342 cm-1 (indicated with an arrow) corresponds to the asymmetric vibration of O-C-O. The spectra shown in (d) and (e) were acquired from a single measurement comprised of 16 scans at 10 kHz and spectral resolution 4 cm-1. The window region of low transmission of the diamond ATR (2200 to 1800 cm-1) has been excluded from both (d) and (e).

with time. The generation of acidic TrisH+ led to the decrease in pH of the liquid (Figure

7.10b). The complementarity data acquisition from the pH and FTIR probes was

+ demonstrated by the ability to simultaneously extract the concentrations of CO2 (aq) and H

+ ions. The real-time characterisation of these two chemical species (CO2 (aq) and H ions)

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could not be followed using a single analytical tool. The molar concentration of CO2 (aq) was calculated using the Beer-Lambert law as:

A = c × l × ε (16) where A is the absorbance, c is the concentration, l is the path length and ε is the molar

6 2 34 extinction coefficient. We used the extinction coefficient for CO2 (aq) (1.5×10 cm /mol) , the path length of the IR light at 2342 cm-1 (0.65 μm)16 and the absorbance data plotted in

Figure 7.10a. Figure 6.10b shows the proton concentration was determined from the variation

+ -pH in pH using [H ] = 10 . The variation in the concentration of CO2 and pH is shown in

Figure 7.10c. After mixing Reagents 1 and 2, the concentration of protons increased (in the

+ form of [TrisH ]) and CO2 (aq) decreased (reaction (14) shifted to the right). From the graph

-1 in Figure 6.10c, the initial rate of consumption of CO2 (aq) was measured to be 0.1 mM s .

A representative ATR-FTIR spectrum of Tris buffer solution (pH = 9.2) is shown in Figure

7.10d. A distinct peak at 1060 cm-1 is observed as a result of C-N stretching. Figure 7.10e shows an ATR-FTIR spectrum of Tris buffer saturated with CO2. The saturation of the Tris

-1 buffer with CO2 resulted in a peak at 2342 cm , which is representative of O-C-O asymmetric stretching. This experiment demonstrated the simultaneous, real-time acquisition of IR spectra and pH data of a flowing solution at a controlled temperature.

7.7 Conclusions

A robust and reliable multi-modal MF analytical platform has been developed that enabled the integration of multiple probes for complementary in situ measurements. The ATR-FTIR, temperature, and pH probes interfaced with a MF reactor enable collection of site-specific real-time information about the chemical state of a reaction and can be used to monitor the

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initial reaction conditions or the change accompanying product generation. Furthermore, we demonstrated the use of temperature measurements to provide feedback to a temperature control unit capable of maintaining on-chip temperatures up to flow rates of 50 mL/h. We showed the utilisation of the integrated probes for the parallel monitoring of multiple variables for various acid/base neutralisation reactions; examination of the relationship between the temperature, pH, and intensity of vibrational absorption bands; and monitoring the kinetics of the reaction between CO2 and a buffer system with clinical applications. This work opens the way for quantitative MF exploration of a large parameter space, which is useful for the optimisation of chemical formulations and determination of their kinetics.

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Chapter 8 Summary, Conclusions and Future Work

8.1 Summary and Conclusions

The overall goal of this thesis was to develop new methods for the studies of chemical and biological phenomena. In particular, this thesis was particularly focused on two distinct areas of research. Chapters 3 and 4 focus on the generation of microenvironments for studies of cell behaviour. The generation of microenvironments was achieved using a high-throughput microfluidic technology for the generation of monodisperse cell-laden microgels. Chapter 6 and

7 describe the development of MF platforms for the study of chemical phenomenon. In chapter 6 we described a novel approach to studying the temperature dependent dissolution of CO2 gas in various liquids. Chapter 7 discusses the development of a modular MF device with integrated analytical probes for real-time, in-situ studies of chemical reactions. The conclusions of each research project are provided in detail below.

In Chapter 3, we presented a novel new strategy for the high-throughput generation of cell- laden agarose microgels. The MF platform provided a fast, efficient, and easy to implement method for the high-throughput generation of cell co-culture libraries which can be analyzed by optical microscopy and flow cytometry. This technique offers speed and control over the total number of encapsulated cells and the relative numbers of different cell populations. We demonstrated the utilization of this approach with an exemplary co-culture (co- encapsulation) system of MBA2 and M07e cells at varying ratios to demonstrate the ability of this approach to modulate paracrine signalling amongst cell populations in a well-defined microenvironment. We further demonstrated the use of this approach as a tool to investigate

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the impact of specific growth factors on a heterogeneous cell population. Co-encapsulation of MBA2 cells with UCB cells was used to determine responsiveness of a sub-population of

UCB cells that are strongly IL-3 dependent.

Chapter 4 described the results of a comparative study of the mechanical properties of the cellular microenvironment on the behaviour of AML2 cancer cells. We studied changes in the mechanical properties (elastic modulus) of macroscopic agarose gels at varying concentration of the biopolymer at room temperature and at 37 oC. We utilized techniques such as rotational atomic force microscopy, fluorescence microscopy, confocal microscopy, and absorbance spectroscopy to study cell behaviour and morphology. Our studies indicated the drastic increase in cell viability and growth in softer matrices (low concentrations of agarose, >2 wt. %) compared with a stiffer matrices (higher concentration of agarose). We previously demonstrated the ability to tune the mechanical properties of agarose microgels in a continuous, high- throughput way.1 In this chapter we demonstrated a MF approach for the high-throughput generation of AML2-laden agarose microgels composed of either 1 or 2 wt. % of the biopolymer.

In Chapter 6 we described an approach to study gas/liquid reactions; specifically the chemical and physical absorbance of CO2 gas in various liquids. We developed a MF method to the temperature-controlled dissolution of gases in liquids that we termed 'bubble breathing', which paved the way for high-throughput studies of temperature mediated CO2 interactions with liquid media. MF technology provides the ability to generate monodisperse bubbles with narrow size distributions. We further demonstrated the ability to monitor the difference in the temperature- controlled dissolution of CO2 in various liquids such as deionized water, ocean water (salt water), and the physically absorbing solvent dimethyl ether of poly(ethylene glycol).

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In Chapter 7, we demonstrated a robust and reliable multi-modal MF analytical platform with multiple integrated probes for complementary in-situ measurements. The probes included an

ATR-FTIR, temperature, and pH probes interfaced directly with the MF reactor. The integration of the probes enabled direct in-situ real-time data acquisition of the chemical state of a reaction, and could be used to monitor the initial reaction conditions or the change accompanying the generation of a product. Furthermore, we demonstrated the use of real-time temperature measurements to provide feedback to a temperature control unit capable of maintaining on-chip temperatures up to flow rates of 50 mL/h. We showed the utilization of the integrated probes for:

(i) the parallel monitoring of multiple variables for various acid/base neutralization reactions; (ii) examination of the relationship between the temperature, pH, and intensity of vibrational absorption bands; and (iii) monitoring the kinetics of the reaction between CO2 and a buffer system with clinical applications.

8.2 Future Work

Encapsulation of cells in microgels is a very promising area of research in polymer materials science, and cell biology; however, several challenges still need to be addressed. First, a large number of cells can be cultured as single cell suspensions, and therefore, there is growing demand for singly encapsulated (isolated) cells in each microgel, without the generation of a large number of empty microgels. Similarly, cell signaling pathways could be more accurately interrogated if the exact number of cells per microgels could be controlled such that co-culture systems could be well defined and easily tuned. New, creative solutions are needed to exceed

Poisson’s distribution in cell encapsulation, without compromising the distribution in microgel dimensions and the reproducibility and robustness of the MF method.

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Second, the formation of libraries of cellular microenvironments often relies on the simultaneous change in several properties of microgels such as microgel size, chemical and mechanical properties, and pore size. Decoupling of the different properties is a significant challenge, which should be addressed through collaborations of cell biologists and polymer scientists. Future work in the design and synthesis of more complex materials will require the use of multi-component systems. The generation of materials from several polymers would enable the ability to tune the properties of the material by compensating for disadvantages of one polymer by the advantages of another polymer.

Third, analytical tools to study encapsulated cells needs to be vastly improved, both in the type of data acquisition, as well as the rate of data acquisition. Analysis of encapsulated cells is primarily performed using optical methods, such as optical microscopy, however, interrogation of cell-laden microenvironments would greatly benefit by high through-put techniques such as flow cytometry.

Microfluidic technology offers an exciting avenue for studying chemical reactions with a

high degree of control over reaction conditions such as stoichiometric ratios of reactants,

reactor temperature, and time of reaction. The use of probes directly integrated into the

MF reactor to monitor in-situ reaction conditions in real-time offers an exciting

opportunity as it allows for direct characterization of the reaction with various analytical

techniques in parallel. In this regard, a highly robust and accurate system for in-depth

investigations of complex gas-liquid systems should yield important information about

the fundamental interactions which are difficult to acquire using conventional bulk

setups. Among the many advantages of MF platforms, segmented flow (two-phase) has

provided a means for high-throughput generation of bubbles for studies of gas-liquid

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reactions. Such systems will be beneficial for future studies of many important

greenhouse gases such as CO2, CH4, NOx, and SO2. In addition, future studies of the

effect of high- and low-molecular weight surfactants on the dissolution of bubbles will

provide insight into the role that organic molecules have on the dissolution of gases in the

oceans. The ability to accurately characterize the early evolution of ultra-fast reactions

will be useful for chemist, biochemists, and engineers. Future work to improve data

acquisition times would be beneficial for increasing temporal resolution for ultra-fast

reactions. Furthermore, further integration of other complex analytical techniques such as

NMR would greatly improve the versatility of MF analytical platforms. Lastly, the

development of MF reactors from more robust materials such as perfluorocarbons (e.g.

Teflon) would allow for the study of more aggressive reactions due to the chemical

inertness of the reactor materials.

References

(1) A. Kumachev, J. Greener, E. Tumarkin, E. Eiser, P. W. Zandstra, E. Kumacheva, Biomaterials 2011, 32, 1477.

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