Digital Microfluidics for ... - University of Toronto

Digital Microfluidics for Multidimensional Biology

Submitted by:

Irwin Adam Eydelnant

MEng, McGill University, 2008

BEng, McGill University, 2007

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Institute of Biomaterials and Biomedical Engineering

University of Toronto

© Copyright by Irwin Adam Eydelnant (2013)

ii

Thesis Title: Digital Microfluidics for Multidimensional Biology

Degree and Year: Doctor of Philosophy, 2013

Name: Irwin Adam Eydelnant

Department: Institute of Biomaterials and Biomedical Engineering

University: University of Toronto

Abstract

Digital microfluidics (DMF) has emerged in the past decade as a novel microfluidic

paradigm. As a liquid handling technology, DMF facilitates the electrostatic manipulation

of discrete nano- and microlitre droplets across open electrode arrays providing the

advantages of single sample addressability, automation, and parallelization. This thesis

presents DMF advances toward improved functionality and compatibility for automated

miniaturized cell culture in two and three dimensions. Through the development and

integration of surface patterning techniques we demonstrate a virtual microwell method

for high precision on-device reagent dispensing in one and two plate DMF geometries.

These methods are shown to be compatible with two-dimensional culture of immortalized

cell lines on ITO, primary cells on coated surfaces, and for co-culture assays. We further

extrapolate this method for the formation of microgels on-demand where form micro scale

hydrogel structures through passive dispensing in a wide array of geometries. With this

system we interrogate three-dimensional cell culture models, specifically for the

recapitulation of kidney epthelialization and the analysis of functional cardiac microgels.

iii

Acknowledgements

A thesis of this nature is produced through the support, guidance, and encouragement of a

community. There are many to whom I am grateful, who I crossed paths over the course of my

PhD tenure, and influenced either myself or my science. As they are too many to list, I

acknowledge a few here who made a particular impact:

Dr. Aaron Wheeler was critical to the success of this thesis, he created a laboratory environment

that knew no barriers for the curious or scientific mind. His unlimited patience allowed for

exploration and learning to occur organically and irrespective of preconceptions.

Dr. William Ryu and his laboratory. They formed my academic home over the course of my PhD

and I’ll always be grateful for being an unofficial part of the group.

Dr. Nathalie Tufenkji for her support and advising during my tenure at McGill University and

the years that followed. She pushed me always to go further and take chances.

My father for his complete support for every decision made at every intersection. For his belief

in the good of people and their ability to affect positive change in this world. His fingers were

crossed for every exam of my academic career including the PhD final defense.

My mother for her belief in my abilities to achieve anything. Her strength and unwavering spirit

have influenced my life greatly. She has always been my biggest fan.

Thank you to you all.

iv

Table of Contents

Acknowledgements ...................................................................................................................................... iii

Table of Contents ......................................................................................................................................... iv

List of Abbreviations .................................................................................................................................. vii

List of Figures ............................................................................................................................................ viii

List of Equations ........................................................................................................................................ xiv

List of Tables .............................................................................................................................................. xv

Overview of chapters ................................................................................................................................. xvi

Overview of author contributions .............................................................................................................. xix

Chapter 1. Introduction ........................................................................................................................... 1

1.1 Historical perspectives on the miniaturization of biology ............................................................ 1

1.2 Microfluidics Paradigms ............................................................................................................... 3

1.3 Digital Microfluidics ..................................................................................................................... 6

1.4 Digital microfluidic theory............................................................................................................ 6

1.4 DMF compatibility with two-dimensional cell culture ............................................................... 14

1.5 Microfluidics for three-dimensional cell culture ......................................................................... 17

1.6 Assays and integration ................................................................................................................ 19

1.7 Future of DMF ............................................................................................................................ 20

Chapter 2. Virtual microwells for digital microfluidic reagent dispensing and cell culture ................. 21

Summary ................................................................................................................................................. 21

2.1 Introduction ................................................................................................................................. 22

2.2 Methods and Materials ................................................................................................................ 25

2.2.1 Reagents .............................................................................................................................. 25

2.2.2 Two-plate DMF bottom-plate fabrication ........................................................................... 25

2.2.3 Two-plate DMF top-plate fabrication ................................................................................. 27

2.2.4 Two-plate DMF device assembly and operation ................................................................ 28

2.2.5 Single-plate DMF device fabrication, assembly, and operation ......................................... 28

2.2.6 DMF dispensing experiments ............................................................................................. 29

2.2.7 Cell Culture and experiments .............................................................................................. 31

2.3 Results and Discussion ............................................................................................................... 31

v

2.3.1 Lift-off patterning ............................................................................................................... 31

2.3.2 Passive dispensing and virtual microwells .......................................................................... 32

2.3.3 Active vs. passive dispensing .............................................................................................. 40

2.3.4 Lift-off vs. protein absorption for passive dispensing......................................................... 41

2.4 Conclusion .................................................................................................................................. 43

Chapter 3. A digital microfluidic platform for primary cell culture and analysis ................................ 44

Summary ................................................................................................................................................. 44

3.1 Introduction ................................................................................................................................. 45

3.2 Methods and Materials ................................................................................................................ 47

3.2.1 Reagents and Materials ....................................................................................................... 47

3.2.2 DMF Device Fabrication and Operation ............................................................................. 47

3.2.3 Primary Cell Isolation and Maintenance ............................................................................. 50

3.2.4 DMF Cell Culture ............................................................................................................... 51

3.2.5 DMF Staining and Microscopy ........................................................................................... 51

3.2.6 DMF Monocyte Adherence Assay ...................................................................................... 52


3.3.1 Digital Microfluidic Primary Cell Culture .......................................................................... 53

3.3.2 Digital Microfluidic Microscopy, Fixation, Permeabilization, and Staining ...................... 56

3.3.3 Digital Microfluidic Monocyte Adhesion Assay ................................................................ 58

3.3.4 Conclusions ......................................................................................................................... 59

Chapter 4. Microgels on-demand ......................................................................................................... 62

Summary ................................................................................................................................................. 62

4.1 Introduction ................................................................................................................................. 63

4.2 Methods....................................................................................................................................... 65

4.2.1 Reagents .............................................................................................................................. 65

4.2.2 DMF device fabrication ...................................................................................................... 65

4.2.3 DMF device assembly and operation .................................................................................. 65

4.2.4 Hydrogel pillar formation and addressing........................................................................... 66

4.2.5 Diffusion analysis and modelling ....................................................................................... 68

4.2.6 Composite hydrogel formation ........................................................................................... 69

4.2.7 Cell Culture ......................................................................................................................... 70

4.2.8 Cell viability and cell distribution ....................................................................................... 70

vi

4.2.9 Epithelialization experiments .............................................................................................. 71

4.2.10 Comparison to conventional fluid handling ........................................................................ 71

4.3 Results and discussion ................................................................................................................ 72

4.3.1 Microgels on-demand ......................................................................................................... 72

4.3.2 Reagent exchange and diffusion into hydrogels ................................................................. 77

4.3.3 Pitch, Multi-Component Arrays, and Composite Microgels............................................... 79

4.3.4 DMF recapitulation of epithelialization .............................................................................. 82

4.3.5 Conclusion .......................................................................................................................... 88

Chapter 5. Cardiac microgels ............................................................................................................... 89

Summary ................................................................................................................................................. 89

5.1 Introduction ................................................................................................................................. 90

5.2 Methods....................................................................................................................................... 92

5.2.1 Reagents .............................................................................................................................. 92

5.2.2 DMF device fabrication ...................................................................................................... 93

5.2.3 DMF device assembly and operation .................................................................................. 93

5.2.4 Cell culture and cardiac microgel formation/addressing..................................................... 94

5.2.5 Cardiomyocyte treatment .................................................................................................... 94

5.2.6 Single cell analysis of cardiomyocyte activity .................................................................... 95

5.2.7 Cardiac activity coefficient (CAC) for fast analysis ........................................................... 95


5.3.1 DMF device fabrication and design .................................................................................... 96

5.3.2 On-demand cardiac microgel formation and assay ............................................................. 97

5.3.3 Higher efficiency analysis of cardiac microgel activity .................................................... 104

5.3.4 Conclusion ........................................................................................................................ 105

Chapter 6. Concluding Remarks and Perspectives on the Future ....................................................... 109

6.1 Conclusions ............................................................................................................................... 109

6.2 Future perspectives ................................................................................................................... 111

References ................................................................................................................................................. 113

APPENDIX A ............................................................................................................................................. 128

vii

List of Abbreviations

2D Two-dimensional

3D Three-dimensional

CAC Cardiac activity coefficient

CM Cardiac myocyte

DMF Digital microfluidics

FITC Fluorescein isothiocyanate

hESC Human embryonic stem cells

FBS Fetal bovine serum

HTS High-throughput screening

ICAM-1 Intercellular adhesion molecule 1

ITO Indium tin oxide

MDCK Marbin Darby canine kidney epithelial cell

MS Mass spectrometry

NBF Neutral buffered formalin

PAEC Porcine aortic endothelial cell

PAVIC Porcine aortic valve interstitial cell

PAVEC Porcine aortic valve endothelial cell

PCB Printed circuit board

PDMS Polydimethylsiloxane

P-S Penicillin and streptomycin

RPMI Roswell Park Memorial Institute medium

SPR Surface plasmon resonance

TCPS Tissue culture polystyrene

THP-1 Human monocytic cell line from acute monocytic leukemia patient

TNF-α Tumor necrosis factor alpha

VCAM Vascular cell adhesion molecule 1

VM Virtual microwell

viii

List of Figures

Figure 1-1: Microfluidic paradigms. (A) Continuous flow channel microfluidic systems can

exploit the laminar flow properties of micron scale confined flows to generate interesting flow

patterns such as the gradient generator depicted here.11

(B) Two phase droplet-in-channel

systems are capable of high-throughput generation of individual droplet compartments. Here two

strategies for droplet formation are depicted within a T-junction (top) and by flow focusing

(bottom).12

(C) Digital microfluidics allows for the manipulation of discrete droplets across

arrays of electrodes. (Permissions requested). ................................................................................ 5

Figure 1-2 Digital microfluidic device geometry. (A) A DMF device consisting of 144

independent actuation electrodes. (B) Devices are composed of two parallel plates. The bottom

plate is patterned with an array of electrodes and coated by a hydrophobic insulator. The top

plate bears the counter electrode and is covered with a hydrophobic coating. (C) Schematic of

droplet translation principles. Separation of charge occurs across the dielectric layer acting on

charges or dipoles in the droplet thereby driving translation. ......................................................... 8

Figure 1-3: Theoretical framework of DMF. (A) Equivalent circuit analysis of DMF driving

force mechanisms. (B) Force estimation for a two-plate DMF device operating on PBS, DI

water, toluene and methanol. Forces are based on a 1 mm2 electrode size, 6 μm of Parylene-C,

235 nm of Teflon- VRMS for a range of

frequencies (100 Hz to 1 MHz). (Adapted from Choi et al.36

– Permissions requested).............. 12

Figure 1-4: Cell culture on DMF. (A) Virtual microwells: Droplets containing cells suspended

in media are translated across patterned hydrophilic sites where a subdroplet is generated by

surface interaction forces. The device is then flipped to allow for cells to settle and adhere to the

hydrophilic site. Here, cells stained with calcein-AM are imaged by stereomicroscopy

immediately after seeding on device. (B) Primary cells: Aortic interstitial cells isolated from pig

hearts cultured for 48 hours on device were then fixed and stained with Hoescht (blue – nuclei)

and Phalloidin (green – actin). Imaging was performed by epifluorescence microscopy. Scale bar

= 200 µm. (C) Multiplexing: Automation combined with multiplexed devices allows for rapid

screening of multiple conditions. Here 16 conditions are screened simultaneously. ................... 16

Figure 2-1: Two-plate digital microfluidic (DMF) device design and assembly. (A) Exploded

view of a device, comprising a bottom plate with patterned electrodes and a top plate bearing

patterned hydrophilic sites. (B) Side-view, not to scale. (C) Schematic depicting two reagent-

dispensing mechanisms on DMF. Active dispensing (i & ii) involves electrostatic stretching of a

reagent from the reservoir followed by splitting. Passive dispensing (iii & iv) occurs

spontaneously as a source droplet is translated across the hydrophilic site. The inset is a three-

dimensional depiction of a virtual microwell, VM (i.e., a droplet formed by passive dispensing).

VM volume is dictated by the diameter of the hydrophilic site (d) and the distance between top

and bottom plates (h). ................................................................................................................... 23

ix

Figure 2-2: X-ray photoelectron spectroscopy evaluation of patterned surfaces. To evaluate the

chemical composition of exposed hydrophilic sites and adjacent Teflon surfaces XPS

measurements were taken on patterned and unpatterned surfaces for comparison: (A) ITO

composition at hydrophilic sites, (B) Teflon on patterned slides, (C) unprocessed ITO surfaces,

and (D) unprocess Teflon surfaces. .............................................................................................. 33

Figure 2-3: Dry passive dispensing to form virtual microwells. (A) Video stills (top-to-bottom)

depicting dry passive dispensing. The dashed circle in panel (i) indicates the position of the

hydrophilic site. (B) Volumes of droplets dispensed in dry passive dispensing as a function of

spacer height and hydrophilic site diameter (n = 5). Asterisks (*) indicate that source droplets

were formed from two actively dispensed droplets. Error bars are 1 S.D. (C) Parameter NVM

calculated for each experimental condition in (B). The shaded region, NVM < 2, indicates

conditions in which two actively dispensed droplets were required to generate the source droplet

for successful passive dispensing.................................................................................................. 35

Figure 2-4: Wet passive dispensing to exchange fluid in a virtual microwell. (A) Video stills

(top-to-bottom) depicting wet passive dispensing in which the virtual microwell contained blue

dye at the hydrophilic site (i) and a red dye source droplet is actuated across the virtual

microwell displacing the blue droplet (ii-v). (B) Multiple passes of reagent across virtual

microwells with varying diameters for 160 µm spacer height. The gray and white bars represent

devices operated with a surrounding matrix of air and mineral oil, respectively. Error bars are 1

S.D. ............................................................................................................................................... 37

Figure 2-5: Single-plate DMF passive dispensing. (A) Picture of a single-plate device depicting

a source droplet and a passively dispensed droplet. (B) Schematic depicting the single-plate

device geometry. ........................................................................................................................... 39

Figure 2-6: Active and passive dispensing as a function of reagent viscosity. Sucrose solutions

of varying viscosity were dispensed on DMF either by active or passive dispensing onto 1500

µm diameter hydrophilic sites (n = 6). Dispensed volumes are plotted as a function of solution

viscosity. Error bars are ± 1 S.D., 95% confidence intervals are indicated by shaded regions, and

the mean dispensed volume for each dispensing mechanism is indicated by a solid horizontal

line................................................................................................................................................. 39

Figure 2-7: Comparison of hydrophilic sites formed by adsorbed protein (on the bottom plate)

vs. liftoff (on the top plate) for dispensing cells into virtual microwells. (A) Results of five trials

seeding MDCK cells (5 105 cells/mL) by passive dispensing. (B) Bright-field images of

MDCK cells seeded on fibronectin coated Teflon and indium tin oxide after 6 hours. Scale bar =

50 µm. ........................................................................................................................................... 42

Figure 3-1: (A) Photograph of DMF device designed for primary cell culture and analysis. A

series of droplets (coloured with red dye for visualization) are positioned at patterned hydrophilic

sites on a device. (B) Schematic of device geometry. The top plate is patterned by a liftoff

procedure to expose hydrophilic sites. The bottom plate bears an array of individually

addressable electrodes with patterned optical windows for imaging. (C) Top and side view

../../../../Desktop/Eydelnant%20-%20Thesis%20v21.doc#_Toc360555494








x

schematic of passive dispensing on hydrophilic sites. (i-ii) A droplet is manipulated to the

hydrophilic site. By actuation of subsequent electrodes the droplet is (iii) stretched then (iv)

passively dispensed, forming a virtual microwell. (D) Side view schematic of device orientation

during experimentation. Devices are maintained right-side up during droplet actuation and are

positioned upside-down during all incubations. ........................................................................... 48

Figure 3-2: Phase contrast images of PAECs, PAVICs, and PAVECs cultured on a DMF device

(top) and in TCPS flasks (bottom). Scale bar = 200 µm. In the DMF images, the bottom plate is

closest to the objective, and the focus is on the layer of cells on the top plate. The cells are

viewed through the circular optical window between two electrodes on the bottom plate (which

are observable but slightly out of focus). ...................................................................................... 57

Figure 3-3: Fluorescent images of PAECs, PAVECs, and PAVICs after fixing, permeabilizing,

and staining on a DMF device. The stains selected for F-actin (FITC-phalloidin, green) and

nuclei (Hoechst, blue). Images were taken at both 10x magnification (top row) and 40x

magnification (bottom row). Scale bar = 200 µm (top row) and 50 µm (bottom row). In these

images, the top plate is closest to the objective. ........................................................................... 57

Figure 3-4: A monocyte adhesion assay performed on DMF-cultured primary PAECs. (A)

Nuclear-stained (Hoechst, red) THP-1 monocytes adhered to PAECs (calcein AM, green).

Representative images of nuclear-stained monocytes adhered to (B) non-stimulated and (C)

TNF-α-stimulated PAECs. In these images, the top plate is closest to the objective. (D)

Monocytes displayed greater adhesion on TNF-α-stimulated PAECs relative to control non-

stimulated PAECs. Data presented as mean ± standard deviation. *P < 0.05. Scale bar = 200 µm.

....................................................................................................................................................... 60

Figure 4-1: Digital microfluidic device geometry. (A) Exploded view of device, comprising a

bottom plate with patterned electrodes and a top plate bearing patterned hydrophilic sites. (B)

Side-view of device, not to scale. (C) Schematic depicting principle of device operation. ......... 67

Figure 4-2: Microgels on-demand. (A-H) Frames from a movie (top) and side-view schematic

(bottom) depicting a sol-state hydrogel droplet containing a fluorescent dye actively dispensed

from a reservoir (A) and then electrostatically manipulated to a patterned hydrophilic site (B &

C). A second droplet is then actively dispensed (D) and passed across the hydrophilic site (E-H).

Scale bars = 2 mm. When the sol-state droplet is passed across the hydrophilic site a sub-droplet

is generated in the shape of the hydrophilic site. Upon crosslinking, each droplet forms a solid

gel pillar. Various geometries can be generated and visualized with epifluorescent

stereomicroscopy (I & J) or confocal microscopy (K & L). A movie depicting the formation of

an array of different microgel shapes can be found online in the supplementary information.

Scale bars = 1 mm. ........................................................................................................................ 73

Figure 4-3: Precision of microgel on-demand formation and reagent exchange. Frames from a

movie (A-D) depicting sol-state Geltrex being passively dispensed to form a sub-droplet. Scale

bar = 2 mm. The diameters of sol-state Geltrex were measured by brightfield microscopy and

compared to the diameters of circular hydrophilic sites (E). The asterisk indicates the single

xi

condition tested for which sub-droplets failed to form. Experiments were performed in triplicate

and error bars indicate ±1 S.D. ..................................................................................................... 75

Figure 4-4: Formation of microgels in varied geometries. Frames from movies depicting the

formation of star (A-D), triangle (E-H), and diamond-shaped microgels (I-L). Microgel density

can be increased by adjusting the pitch of patterned hydrophilic sites on the device top-plate (M).

A movie depicting the formation of an array of microgels in different shapes can be found in the

online supplementary information. Scale bars = 2 mm. ............................................................... 76

Figure 4-5: Off-chip microgel analysis. Agarose (5% w/v) hydrogels were formed on device

then allowed to crosslink at ambient temperature. Top-plates were removed to expose the

hydrogel allowing for imaging by environmental scanning electron microscopy. ....................... 78

Figure 4-6: Reagent exchange in hydrogel pillars. (A-D) Frames from a movie depicting a

droplet of fluorescein being electrostatically manipulated across a (transparent) microgel pillar

and subsequent diffusion of fluorescein into the microgel. Scale bars = 2 mm. (E) Experimental

(red dots), empirical fit (dashed blue line), and simulation (solid black line) of diffusion profiles

for fluorescein into Geltrex microgels. The inset shows a fluoresecent image (left) and heat-map

simulation of concentration (right) of fluorescein at the half-saturation point 1/2). (F) Apparent

diffusion (Da) coefficients measured for fluorescein, 4 kDa FITC-dextran, and 40 kDa FITC-

Dextran into Geltrex. Error bars indicate 1 S.D. ........................................................................... 80

Figure 4-7: Precision of reagent exchange across microgels. Frames from a movie (A-D)

depicting a droplet of reagent passed across a microgel. The diameters of dispensed droplets

were measured by brightfield microscopy and were compared to the diameters of circular

hydrophilic sites (E). The asterisk indicates the single condition tested for which sub-droplet

failed to form. Experiments were performed in triplicate and error bars indicate ±1 S.D. .......... 81

Figure 4-8: Pitch, multicomponent arrays, and composite microgels. 32-plex arrays of microgels

were formed from Geltrex supplemented with fluorescein for visualization (A). Combinatorial

arrays of Geltrex microgels were formed on device containing mixtures of red, yellow, and/or

green microspheres (B). Gradient bars indicate the percent compositions of each respective

microsphere, from 100% red at the left and 100% green at the right. Composite microgels

containing fluorescent microspheres were formed with inner agarose and outer Geltrex layers (C:

green filter, D: red filter, E: composite image). Scale bars = 2 mm. ............................................ 83

Figure 4-9: Higher order tissue formation and handling in microgel pillars. Spheroids are

generated five-plex in microgels on DMF (A). The inset highlights one microgel containing

spheroids. Confocal microscopy image stack of MDCK cells in a Geltrex pillar demonstrates cell

distribution throughout the z-axis (B). Images depicting MDCK spheroid formation in a gel pillar

over four days in culture (C-F). Cells were stained for actin with phalloidin (green) and nuclei

(blue). On day four lumen formation was observed. Hydrogels bearing 4-day MDCK spheroids

were subjected to media exchange five consecutive times either by automated pipetting or DMF.

Representative bright-field images of spheroids pre- and post-manipulation for both systems (G-

J). Scale bars = 20 µm. Graph of spheroid deformation for five trials evaluating 10 spheroids

xii

each using robotic pipetting into a multiwell plate or DMF (K). The deformation value of each

spheroid is indicated by a blue (DMF) or red (conventional reagent delivery) dot with black bars

representing the mean deformation of each trial. Bar graphs show overall deformation averages

across the five trials for each respective system. Error bars show 1 S.D. In three of the trials,

identified by asterisks, the conventional method dislodged the hydrogels from the well, resulting

in material failure. ......................................................................................................................... 85

Figure 4-10: Z-axis distribution of cells in microgels. Graph of distribution of cells in the z-axis

for the data in Figure 5B in the main text. The number of cells was enumerated at each image

plane (~6 µm thick). ...................................................................................................................... 86

Figure 4-11: Spheroid size. Comparison of spheroid size for three-dimensional MDCK culture in

microscale (DMF, white bars) and macroscale (96-well plate, gray bars). .................................. 86

Figure 5-1: Cardiac microgel formation on DMF. A sol-phase hydrogel droplet containing a

suspension of cardiac myocytes is drawn from a reservoir (A). The droplet is translated across

hydrophilic sites that are positioned (on the transparent top plate) above the optical windows on

the bottom plate (B-F). The white arrows indicate the direction of droplet movement. Scale bar =

4 mm. ............................................................................................................................................ 98

Figure 5-2: Cell distributions in microgels as a function of seeding density. Phase contrast

images of micorgels formed with cells seeded at low (A), moderate (B), or high (C) density. The

cells are visible through windows on the bottom plate of the device. At low and modest densities,

cells are homogenously dispersed through the hydrogel. At high density, cell aggregation is

apparent. Scale bar = 250 µm. .................................................................................................... 100

Figure 5-3: Brightfield microscopy of CM microtissues. Images of CMs in a microgel formed

on-demand at 4x (A), 10x (B) and 20x (C) magnification. Images of cells in the same microgel at

........................... 100

Figure 5-4: Confocal images of cardiac microgel on DMF. A cardiac microgel was fixed and

stained (blue for nuclei and red for actin) after five days of culture on DMF. Imaging was

performed at 5× (A), 10× (B), and 20× (C) magnification, with elongated branched cells evident

at each level. Scale bars = 100 µm, 50 µm, and 25 µm respectively. ......................................... 101

Figure 5-5: 3D cross-section of cardiac microgel. Cells were fixed and stained on DMF (blue

for nuclei, red for actin). Maximum combined exposure of an orthographic view in (A) z-x, (B)

x-y, and (C) y-z planes, demonstrates a well-connected network of CMs that are distributed in

3D space. Scale bar = 50 µm. ..................................................................................................... 101

Figure 5-6: Cardiomyocyte function in microgels. Cardiac microtissues were cultured for four

days on DMF and evaluated before and after exposure to epinephrine. The areas of twenty

individual cells were calculated and plotted with respect to time – representative traces (blue-

unstimulated; red – EPI-stimulated) for one cell are shown in (A). Insets microscopy image

demonstrates the contraction measurements performed on single cells. Black line indicates shape

pre-contraction, yellow line indicates shape post-contraction. These data were pooled and

evaluated for contraction frequency (B), contraction distance (C), and duration (D) for control

xiii

and epinephrine stimulated cells. Experiments performed in triplicate and error bars indicate ±1

S.D. ............................................................................................................................................. 103

Figure 5-7: Cardiomyocyte activity coefficient. After five days of culture on DMF, a beating

cardiac microgel was treated with a fixative solution and observed at 10× magnification for 30

minutes (A). A heat map was generated to represent cumulative cardiomyocyte activity

coefficients at 1 (B), 15 (C), and 30 min (D) after exposure to fixative. Heat maps are presented

as overlays on phase contrast microscopic images. Here blue indicates regions of low activity

and red indicates high activity. With increased exposure to fixative over time, reductions of

activity are observed (as shown through reduced red regions in the heat maps). ....................... 107

Figure 5-8: Cardiac activity coefficient of stimulated cells. CACs were calculated for control

and epinephrine-treated cardiac microgels, and the response (A) was significantly different by a

one-tailed t-test (p < 0.01). Heat maps representing cardiomyocyte activity generated for control

and epinephrine-stimulated microgels (B,C respectively), and brightfield images used for

analysis (D, E respectively). In these images red regions indicate higher activity, with blue

regions indicating lower activity. Scale bars = 50 µm. ............................................................... 108














xiv

List of Equations Eqn. 1.1 ........................................................................................................................................... 3

Eqn. 1.2 ........................................................................................................................................... 7

Eqn. 1.3 ......................................................................................................................................... 10

Eqn. 1.4 ......................................................................................................................................... 10

Eqn. 1.5 ......................................................................................................................................... 10

Eqn. 1.6 ......................................................................................................................................... 10

Eqn. 1.7 ......................................................................................................................................... 10

Eqn. 1.8 ......................................................................................................................................... 10

Eqn. 1.9 ......................................................................................................................................... 10

Eqn. 1.10 ....................................................................................................................................... 11

Eqn. 1.11 ....................................................................................................................................... 11

Eqn. 1.12 ....................................................................................................................................... 11

Eqn. 1.13 ....................................................................................................................................... 11

Eqn. 1.14 ....................................................................................................................................... 13

Eqn. 1.15 ....................................................................................................................................... 13

Eqn. 2.1 ......................................................................................................................................... 36

Eqn. 4.1 ......................................................................................................................................... 69

Eqn. 4.2 ......................................................................................................................................... 69

Eqn. 4.3 ......................................................................................................................................... 69

Eqn. 4.4 ......................................................................................................................................... 72

Eqn. 5.1 ......................................................................................................................................... 95

Eqn. 5.2 ......................................................................................................................................... 96

xv

List of Tables

Table 1-1: Comparison of well-plate and microfluidic methods used for cell culture .................. 5

Table 2-1: Elemental identification and quantification of species detected on patterned DMF

surfaces using XPS. Samples labels refer to Figure 2-2. .............................................................. 33

Table 3-1: Comparison of adherent cell culture using DMF between the new methods reported

here and those previously published. ............................................................................................ 61

xvi

Overview of chapters

This thesis is a comprehensive summary of projects related to the development of a digital

microfluidic platform for the culture and analysis of primary cells in two and three dimensional

cell culture.

Chapter one provides an overview of microfluidics in biological applications with an emphasis

on digital microfluidic platforms and theory. Recent developments within the field that have

enabled the implementation of DMF for cell culture and assay are reviewed.

Sections of this introduction were published in the article: Eydelnant, IA, and Wheeler, AR.

Digital microfluidic cell culture. BioTech International. (2012), 24, 20-22.

Chapter two describes the development of ‘virtual microwells’ (VMs) for reagent dispensing and

cell culture on DMF. The hydrophilic patterning technique developed here serves as the basis for

much of the work described in subsequent chapters within this thesis. With this method we were

able to reliably dispense volumes from 80 to 800 nL in air and oil, improve cell culture on DMF,

and perform the first example of reagent dispensing on a single plate DMF device. Further we

demonstrated the first example of passive dispensing in oil filled devices. A quantitative criterion

was also formulated to guide the design of such hydrophilic sites in future DMF systems. The

methods developed in this work are already finding applications in many DMF and non-DMF

applications.

This work resulted in the following publication: Eydelnant IA, Uddayasankar U, Li BB, Liao

MW, Wheeler AR. (2011) Virtual microwells for digital microfluidics. Lab on a Chip. 12, 750-

757.

Chapter three describes the first digital microfluidic method for the culture and analysis of

primary cells. Utilizing the virtual microwell method developed in chapter two, primary porcine

aortic endothelial, valve endothelial, and valve interstitial cells were cultured, fixed, and stained

xvii

in microlitre droplets using electrostatic manipulation of reagents. A monocyte adhesion assay

was performed completely on device supporting the implementation of DMF for cell culture and

assay. This study reinforces the applicability of DMF for the culture of sensitive cell types in

sub-microlitre droplets and looks toward integration of DMF with physiologically relevant

models of biology.

This work resulted in the following publication: Srigunapalan S*, Eydelnant IA*, Simmons C,

Wheeler AR. (2011) A digital microfluidic platform for primary cell culture and analysis. Lab on

a Chip. 12, 369-375. *Equal contribution

And the following proceedings: Eydelnant IA, Li BYB, Chang WC, Stanford W, Wheeler AR.

(2011) Upside-down digital microfluidic based embryonic stem cell culture. Proc. Micro. Tot.

Anal. Sys. (33-35)

Chapter four describes a novel technique for microscale hydrogel formation termed ‘microgels

on-demand’. This work builds on the virtual microwells presented in chapter two and the

determination of biocompatibility of DMF for cell culture in chapter three. This is the first

method capable of generating microscale thermally cross-linked hydrogels in a range of

geometries. Until now, this has only been possible with photo-crosslinking chemistries. Further,

we implement this system for three-dimensional cell culture, specifically to recapitulate a model

of kidney epithelialization. We demonstrate the utility of fully automated DMF operation for the

generation of high density microgel arrays, the formation of combinatorial microgels on-device,

and composite microgels consisting of multiple hydrogel constituents.

This work is the subject of a manuscript that has been submitted for publication: Eydelnant IA,

Li BB, Wheeler AR. (2013) Microgels on-demand (Submitted).

And the following proceedings: Eydelnant IA, Li BB, Wheeler AR. (2012) Virtual microwells

for three-dimensional cell culture on a digital microfluidic platform. Proceedings of the IEEE

25th International Conference on Micro Electro Mechanical Systems (MEMS). 898-901.

xviii

Chapter five presents the implementation of microgels on demand, developed in chapter 4, for

functional drug screening of cardiac microgels. Cardiomyocytes were seeded in sub-microlitre

microgels and cultured on device for up to one week. After three days of culture spontaneous

contractions were observed within the construct and coordinated contractions became evident at

five days after seeding. We perform single cell analysis to determine contraction frequency,

magnitude, and duration with the finding that these constructs are functional and respond to drug

treatments in accordance with previous studies in the literature. To improve throughput of

analysis, an image processing method was developed to analyze broader construct activity

resulting in a metric referred to as the cardiomyocyte activity coefficient (CAC). We present

preliminary evidence that this metric can be correlated to the contraction frequency.

This work is the subject of a manuscript that is currently in preparation: Eydelnant IA,

Thavandiran N, Radisic M, Wheeler AR. (2013) Cardiac microgels (In preparation).

And the following proceedings: Eydelnant IA, Li BB, Wheeler AR. (2012). Digital

microfluidics for on-demand 3D microgel formation and functional myocardial tissue assays.

(2012). Proc. Micro. Tot. Anal. Sys. (112-114).

xix

Overview of author contributions

Chapter two describes the implementation of virtual microwells for DMF. This project was

initiated, designed, and lead by me. Fellow graduate student Uvaraj Uddayasankar provided

helpful discussions that initiated this work. I developed the Teflon liftoff protocol, fabricated

devices, performed dispensing and cell experiments, and developed the non-dimensional number

reported. Undergraduate researchers Bingyu ‘Betty’ Li and Donna Liao reproduced the

dispensing experiments for all conditions, assisted with device fabrication, and prepared reagents

for the experiments.

Chapter three demonstrates culture and analysis of primary cells for the first time on DMF.

These projects resulted in a co-first authored paper shared between graduate student Suthan

Srigunapalan (SS). I designed and fabricated devices then tested reagent compatibility. For the

primary cell work, cells were cultured by SS. Together SS and I performed all experiments for

cell seeding, fixing, staining, and functional analysis. SS imaged the cells and did the analysis for

the adhesion assay.

Chapter four describes a method we term `microgels on-demand`. For this work I selected

appropriate hydrogel materials, tested compatibility with surfactants, assessed the capability of

using virtual microwells to form microgel structures, designed geometries, performed electron

scanning microscopy imaging, confocal imaging, cell seeding, and analaysis. Bingyu `Betty` Li

assisted with kidney epithelialization experiments and was instrumental in the selection of

fixative for this work. She also performed replicate studies on volume dispensing in virtual

microwells.

Chapter five describes the implementation of microgels on demand for cardiac microgels. For

this work I initiated the collaboration with the Radisic laboratory. I seeded cells on device,

cultured them, ran functional assays, and analysis. Both the single cell analysis and

xx

cardiomyocyte activity coefficient were developed by myself. Nimalan Thavandiran (then

graduate student) provided helpful discussions, neonatal cardiomyocytes and media for

experiments.

1

Chapter 1. Introduction

1.1 Historical perspectives on the miniaturization of biology

Prescient scientists in the mid-20th

century set the stage for a future where accessing biology at

the micron scale would become commonplace. In 1944 Schrodinger asked the famous question

‘What is life?’ at his lecture at the Dublin Institute for Advanced Studies, peering into the nano-

and micron scale components of the cell from a purely physical and chemical perspective.1

Feynman inspired with his visionary and speculative 1960 lecture ‘There’s Plenty of Room at the

Bottom’ introducing the untapped field of miniaturization for electronics and biology.2

Concurrently, significant thought was being given to Brownian motion and diffusion by

Einstein3 and Berg

4 in biological systems while Taylor

5 had already begun examining liquid

flows through micron scale channels. The ability to implement these concepts in biological

investigation through microfluidic systems gained significant ground with the development of

miniaturized chromatographics5,6

and ink jet technologies.7 These methods remained inaccessible

to the majority of scientists due to requisite specialized equipment, borrowed from the semi-

conductor industry, which permitted channels to be etched in glass and silicon. With the

development of soft-lithography and subsequent enabling of replica moulding, in cheap

elastomers such as polydimethylsiloxane (PDMS), pioneered by the Whitesides group in 1998,8

the field of microfluidics experienced an immediate explosion of accessibility, spawning

microfluidic applications in nearly every imaginable scientific discipline.

2

Parallel to early works in microfluidics and movement toward miniaturization, bench-top biology

was undergoing rapid change, initiated by Hungarian physician Takatsy in 1955 when he

proposed a move away from traditional glass tubes and dishes by developing the 96-well plate to

increase laboratory throughput.9 The microtitre plate provided a platform for early bench-top

miniaturization which today is driven by the high-throughput demands of contemporary drug

screening and systems approaches to biology. High-throughput screening (HTS) emerged in the

1980s and 1990s as a new standard for drug discovery within the pharmaceutical industry, where

microtitre formats increased to 384 and 1536 wells, thereby maximizing experimental densities

within equivalent footprints and further decreasing reagent consumption. Since then, the

technologies that emerged from this field have spilled over extensively into broad fields of study

including genomics, protein crystallization, materials science, and environmental toxicity

sampling.

Today HTS methods have reached a standstill – throughput with current automated liquid

handling robotics (ALHR) and microtitre plates has plateaued and more attention is now being

paid toward higher content and quality of data. The adoption of high-throughput cell-based

biological screens as standard practice within academic laboratories is furthering the need for

accessible high-throughput systems. Robotic liquid handling – though useful for experimental

automation – is often inaccessible because of high capital costs (e.g., robotics), large volumes of

reagent consumption (e.g., drugs, media, cells) and high turnover of consumables (e.g.,

microwell plates, pipette tips). Data generated in standard HTS assays does not typically include

sufficient content to instruct lead pursuit without further experiments. These obstacles have

motivated the development of miniaturized platforms with the capability to manipulate micro-

3

scale samples. Microfluidic technologies, where liquid manipulation is implemented in micron-

scale confined volumes, potentiate the development of the tools needed for ultra-low volume

reagent handling. With an abundance of research groups developing biologically oriented

microfluidic technologies, the first generation of laboratory protocols implemented in devices

with the footprint of a credit card has been realized enabling improved control over experimental

conditions and higher density experimental footprints.

1.2 Microfluidics Paradigms

Multiple paradigms of microfluidics, including continuous channel microfluidics, droplet-in-

channel microfluidics, and digital microfluidics, have emerged, each with its own advantages

and disadvantages (Table 1-1).10

Continuous channel microfluidics, where reagent flows are

confined within micron scale channels, allow for well controlled serial device operations. Flow

properties of these systems are readily described by the dimensionless Reynold`s number:

vL

ForcesViscous

ForcesInertialRe Eqn. 1.1

where ρ is the fluid density (kg/m3), v is the mean velocity (m/s), L is the characteristic length of

the system, and υ is the kinematic viscosity (m2/s). Flow in channel microfluidic systems is

characterized by viscous dominated flow and low Reynold’s number regimes, where multiple

flow streams can be constrained within a single channel without mixing. These features have

been exploited in biological assays for the formation of well-defined diffusion gradients across

cell monolayers, single-cell capture and analysis (e.g., PCR, fluorescence microscopy), modeling

4

microvasculature, and other studies requiring precise control of the chemical (e.g., growth

factors, cytokines) and physical (e.g., shear stress, flow rate) cellular microenvironment. With

the incorporation of on-device valves, these devices have been scaled into compartmentalized

high-throughput systems. However, because of the inherently complex fabrication protocols

required to form such systems, reliance on external equipment (e.g., pumps, valve manifolds),

and networks of connecting tubing required for reagent transfer, biologists have been slow to

adopt such systems into their routine work-flow.

In droplet-in-channel microfluidics, pico- and nano-litre droplets are generated in a two-phase

flow in microchannels. This allows for the formation of thousands of independent droplets per

second that can be merged, sorted, and reacted. With simple fabrication and high-throughput

operation (10-100 kHz), these devices are well positioned for screening in biological

experiments. Biocompatible surfactants stabilize the emulsions, allowing for encapsulation of

live cells for suspension culture or hydrogel materials for adherent culture. The use of fluorinated

oils in the continuous phase permits sufficient oxygen transfer to maintain cell viability for

multiple days. The throughput of droplet microfluidic systems is unparalleled; however, such

systems are not well suited to multi-step long-term applications involving cells because of the

challenges in addressing individual droplets (for media exchange, reagent addition, staining). In

addition, both continuous and droplet-in-channel methods typically require dedicated specialized

microscopy analysis techniques; this limits their flexibility for integration with standard

analytical laboratory equipment. Over the past decade, a third paradigm, digital microfluidics,

has emerged as a potential solution to these limitations.

5

Figure 1-1: Microfluidic paradigms. (A) Continuous flow channel microfluidic systems can

exploit the laminar flow properties of micron scale confined flows to generate interesting flow

patterns such as the gradient generator depicted here.11

(B) Two phase droplet-in-channel

systems are capable of high-throughput generation of individual droplet compartments. Here two

strategies for droplet formation are depicted within a T-junction (top) and by flow focusing

(bottom).12

(C) Digital microfluidics allows for the manipulation of discrete droplets across

arrays of electrodes.

Table 1-1: Comparison of well-plate and microfluidic methods used for cell culture

Criteria Well-plate Microchannel

Droplets in

Channels

Digital

Microflfuidics

Cost $1-2 per plate <$10 / PDMS

device

<$10 / PDMS

device $10-50 / device

Reagent volumes µL-mL nL- µL pL-nL nL-mL

Throughput High Moderate High Moderate

Automation Yes Yes Yes Yes

No. of scientists >100,000 >1,000 <500 <20

6

1.3 Digital Microfluidics

Digital microfluidics (or DMF) is a liquid handling technology that permits the independent

electrostatic manipulation of individual pico- to micro-litre size droplets across arrays of

electrodes.13

The most common DMF format features droplets sandwiched between two plates,

and typical operations include droplet dispensing, splitting, merging, and mixing.14

The bottom

plate is patterned with electrodes buried beneath a hydrophobic insulator. The top plate

comprises a contiguous electrode coated with a hydrophobic layer. The hydrophobic coatings are

critical to reduce friction forces that can impede droplet movement. When voltages are applied to

a driving electrode on the bottom plate relative to a counter-electrode on the top plate, a

separation of charge occurs across the insulator acting on ions or dipoles within the droplet. The

resulting electrostatic force drives droplet translation. These devices are commonly operated in

air, though filler media such as silicone oil can be used to reduce voltages needed to move

droplets. Device fabrication follows basic photolithography and metal etching protocols on a

range of substrates including glass, silicone, flexible polyimide films, compact discs, and printed

circuit boards (PCBs). Currently a range of techniques for rapid prototyping and mass-

production of devices are being investigated, with an emphasis on multi-layer PCBs15

and high

density thin film transistor (TFT) arrays.16

1.4 Digital microfluidic theory

The physics of DMF droplet actuation has not been completely elucidated, with multiple

mechanisms described in the literature. The forces controlling droplet motion can be divided into

driving and resistive forces. Historical understanding of DMF associated driving forces often

begin with Pellat’s17

original observation of the generic phenomenon related to what we now call

7

DMF over a century ago where he demonstrated that an external voltage can cause an insulating

dielectric liquid to raise upward against gravity when confined between vertical parallel

electrodes. In some instances this phenomenon is described as electrowetting or electrowetting

on dielectric (in the case where electrodes are coated with dielectric material, this is often

described with the acronym, EWOD)18–26

. This follows from the observation that upon

application of a potential across the electrode, droplets of fluid with high surface tension (e.g.,

water) tend to wet the surface (i.e., experience a significant decrease in contact angle) that can be

described within a thermodynamic context by the Young-Lipmann relation27

:

d

V

2coscos

2

r00 Eqn. 1.2

where θ and θo are the static contact angles with and without applied voltage respective, εo is the

permittivity of free space, εr is the relative permittivity of the dielectric, V is the applied voltage,

and γ is the air-liquid surface tension. Here droplet movement is assumed to occur due to

capillary pressure that results from asymmetric contact angles across the droplet. However,

EWOD theory fails to describe two key observations: (1) low surface tension liquids that have no

apparent change in contact angle are movable on such systems28

(and indeed, have been used by

8

Figure 1-2 Digital microfluidic device geometry. (A) A DMF device consisting of 144

independent actuation electrodes. (B) Devices are composed of two parallel plates. The bottom

plate is patterned with an array of electrodes and coated by a hybdrophobic insulator. The top

plate bears the counter electrode and is covered with a hydrophobic coating. (C) Schematic of

droplet translation principles. Separation of charge occurs across the dielectric layer acting on

charges or dipoles in the droplet thereby driving translation.

9

the Wheeler group for many applications), and (2) θE cannot be reduced to zero upon application

of infinitely high voltage (as suggested by the Young-Lippman equation), and in fact saturates at

some intermediate value determined by the particular liquid and dielectric solid being used.

Kang29

proposes that this contact saturation arises from the free-energy contribution of the

electrical double layer in the liquid phase, which would be incapable of driving the observed

changes in contact angle. Buehrle and Mugele30

compute the liquid profile near the contact line

by enforcing electromechanical equilibrium at the interface while minimizing energy. Their

result further contradicts EWOD theory as there exists only an ‘apparent’ change in contact angle

since at a given distance from the dielectric surface the contact angle asymptotically approaches

θo. Jones31

furthers this argument through an electromechanical approach demonstrating contact

angle change as a consequence of a strong electric field near the three-phase contact line. In his

work he proposes a Gedanken experiment modelled after the height of rise experiment described

above, however in his example, he restrains the meniscus shape (and effectively contact angle)

by floating a thin membrane at the top of the column while retaining the height of rise with the

application of a voltage to the electrodes.

Given the uncertainties surrounding the EWOD mechanism, we and others propose that a more

complete illustration of droplet actuation comes from electrodynamic analysis29,31–33

. Under this

treatment droplet movement is explained by the action of electric forces on free charges in the

droplet meniscus of conductive liquids or dipoles inside the droplet of dielectric liquids. As such

this model overcomes the limitations of EWOD demonstrating droplet movement without

requiring wetting and rationalizes contact angle saturation as a balance between electrical and

10

surface tension forces. The theoretical content of electrodynamics is contained within the

Maxwell equations and Lorentz’s Force Law34

:

Gauss’s Law: o

E 1 Eqn. 1.3

Unnamed: 0 B Eqn. 1.4

Faraday’s Law tBE

Eqn. 1.5

Ampere’s Law: tEJB ooo Eqn. 1.6

Lorentz’s Law: )( BEqF Eqn. 1.7

The total electromagnetic force on the charges in a volume V can be determined by Lorentz’s

force law that describes the effects of electromagnetic fields on charges:

VV

BJEBEF )()( Eqn. 1.8

Rewriting this equation as a force per unit volume and substituting Maxwell’s equations (Eqn.

1.3) and (Eqn. 1.6) to eliminate ρ and J, leaving only the field terms we find:

Bt

EBEEf oo

)1()(

Eqn. 1.9

where the first term accounts for Coulombic forces and the second term magnetic forces.

Further manipulation brings this expression to the form:

11

)()1(2

1)()(1)()( 22 BEt

BEBBBBEEEEf oo

oo

o

Eqn. 1.10

Neglecting the magnetic field components, and reverting to the volumetric integral we find:

V

dEEEEEF )(5.0)()( 2 Eqn. 1.11

Simplifying the above volumetric integral to a surface integral by the introduction of the

Maxwell stress tensor facilitates solution described previously for DMF droplet

manipulation29,32,35

:

S

ndsTF Eqn. 1.12

)2

1( 2EEET ijjiij Eqn. 1.13

Here Tij is the stress tensor, where i and j refer to pairs of x, y, and z axes, δij is the Kronecker

delta function, and E is the electric field surrounding the droplet. In this electrostatic

interpretation, Tij is the stress (i.e., force per unit area) of the ith direction on an element of

surface oriented in the jth direction. Here, diagonal elements represent pressures and off-

diagonal elements are shears. The Wheeler group recently used this electrodynamic model to

estimate the driving forces on droplets of water in eight different DMF device geometries, and

the results correlated well with experimental observations32

Equivalent results can be determined through a circuit representation of the DMF device (Figure

1-3A) within an electromechanical framework.31

In this analysis the amount of energy,

12

Figure 1-3: Theoretical framework of DMF. (A) Equivalent circuit analysis of DMF driving

force mechanisms. (B) Force estimation for a two-plate DMF device operating on PBS, DI

water, toluene and methanol. Forces are based on a 1 mm2 electrode size, 6 μm of Parylene-C,

235 nm of Teflon-AF, a gap size of 150 m and an applied voltage of 100 VRMS for a range of

frequencies (100 Hz to 1 MHz). (Adapted from Choi et al.36

)

13

E, stored within the system can be calculated as a function of the applied voltage frequency and

droplet position along the direction of translation. This method assumes that the cross-sectional

area of the drop is readily approximated by a square with sides length L:

i i d

fjVxL

d

fjVx

LxfE

i

filleri,fillerr.i,

i

liquidi,liquidr.i, )2()2(

2),(

Eqn. 1.14

where εr,i,liquid, Vi,liquid, and εr,i,filler, Vi,filler are the relative permittivity and voltage drop for the

liquid and filler fluid portion of the electrode respectively, and di is the thickness of layer i

(corresponding to the dielectric, hydrophobic, liquid, or filler layers). The associated change in

energy as x progresses from 0 to L is equivalent to the work done on the system. Differentiation

of Eqn. 1.12 with respect to x yields the driving force as a function of frequency:

i i d

fjV

d

fjVL

x

xfEfF

i

filleri,fillerr.i,0

i

liquidi,liquidr.i,0 )2()2(

2

),(

Eqn. 1.15

Typical DMF droplet manipulation employs AC frequencies on the order of kHz. At these

frequencies, the majority of the voltage drop occurs across the dielectric layer. Estimates of

forces by Eqn. 1.15 indicate magnitudes in the range of µN (Figure 1-3B). Above a critical

frequency, fc, the electric field shifts from the dielectric to the droplet resulting in a liquid-

dielectrophoretic regime (Figure 1-3B). For these reasons liquids with low conductivity and

permittivity require prohibitively high voltages, which can be remedied through mixing with

reagents with more amenable properties.

14

Droplet movement on DMF must be considered as a balance of electrostatic forces (described

above) and resistive forces. The major resistive forces are friction between the droplet and the

hydrophobic surface, contact line pinning, and viscous drag.22,37–39

The first are determined by

the nano- and microscale roughness of the hydrophobic surface, the second is a molecular

adhesion effect occurring at the three-phase (surface, liquid, air) contact line of a droplet that

results in ‘sticking’ thereby impeding droplet movement, while the third are associated directly

with displacement of filler fluid during droplet translation. Contact line pinning is implicated in

DMF observations of contact angle hysteresis and neglecting its effects results in inaccurate

simulations of droplet motion. Modeling this effect is challenging as simulations require

nanometer length scales and nanosecond time scales. For a droplet to move on DMF the electric

field generated across the dielectric must be strong enough to overcome these resistive forces and

can require applied voltages from 10 Vpp to 1000 Vpp depending on dielectric coatings and the

liquid being manipulated.

1.4 DMF compatibility with two-dimensional cell culture

Digital microfluidics is a useful platform for the miniaturization of cell culture and assays. With

the capacity to support both adherent and suspension cell culture, several research groups have

demonstrated long-term culture and passaging on-device. Suspension cell culture is well suited

to DMF as droplets are manipulated across hydrophobic surfaces that are resistant to adhesion.

DMF based viability assays have been performed with comparable results relative to 96-well

plate assays and 100-fold reductions in reagent volumes.40

Droplet mixing by translation allows

for cell growth, and droplet splitting and merging are useful for dilution and passaging. For

adherent cell culture, multiple strategies have been developed for surface functionalization

15

including protein deposition, plasma etching, and a fluorocarbon liftoff technique described in

Chapter 2 of this thesis. Each has been demonstrated for the culture of immortalized cell lines,

while the latter has been successfully implemented in the culture of more sensitive cell types

including primary cells as presented in Chapter 3 (Figure 1-4A & B).41

The introduction of

hydrophilic sites to the hydrophobic coating resulted in the discovery of a novel fluidic

phenomenon termed passive dispensing. As droplets are translated across hydrophilic sites, a

portion of the droplet is pinned to the site and a sub-droplet is formed. Chapter 3 examines

passive dispensing for controlling droplet volumes in cell seeding and subsequent media and

reagent exchange.42

The successful culture of multiple cell types suggests minimal electromagnetic effects on DMF

cultured cells. This has been supported by computational modeling of potentials across the

device, which demonstrated that the majority of the voltage drop occurs in the dielectric layer

and a minimal potential is experienced by the droplet. Recent microarray analysis of cell lines

actuated in DMF-like devices indicate minimal differences in transcriptional profiles at normal

operating conditions.43

Biofouling of device surfaces remains the most significant challenge in

long-term device operation. Protein adsorption to the hydrophobic surfaces results in droplet

pinning thereby restricting droplet translation. The addition of low concentrations of

biocompatible surfactants (e.g., Pluronics) improves device function, but does not allow for

indefinite device operation. Novel surfactants and fouling resistant surface coatings are being

investigated to address these issues.

16

Figure 1-4: Cell culture on DMF. (A) Virtual microwells: Droplets containing cells suspended

in media are translated across patterned hydrophilic sites where a subdroplet is generated by

surface interaction forces. The device is then flipped to allow for cells to settle and adhere to the

hydrophilic site. Here, cells stained with calcein-AM are imaged by stereomicroscopy

immediately after seeding on device. (B) Primary cells: Aortic interstitial cells isolated from pig

hearts cultured for 48 hours on device were then fixed and stained with Hoescht (blue – nuclei)

and Phalloidin (green – actin). Imaging was performed by epifluorescence microscopy. Scale bar

= 200 µm. (C) Multiplexing: Automation combined with multiplexed devices allows for rapid

screening of multiple conditions. Here 16 conditions are screened simultaneously.

17

1.5 Microfluidics for three-dimensional cell culture

Hydrogel based three-dimensional (3D) cell culture is rapidly becoming a fundamentally

important tool in biological research.44

Hydrogel materials can be derived from inert or animal

sources providing for customizable microenvironments to elucidate or direct cell function and

behaviour. Bissell and coworkers45

were the first to demonstrate that in certain models of

epithelialization, cells traditionally cultured in monolayers would form polarized hollow

spheroid structures when dispersed in collagen hydrogel matrices. Microarray analyses of cells

isolated from these spheroid structures confirm that they are transcriptionally more

representative of in vivo conditions than two-dimensional (2D) culture.46

This transition from

monolayer to 3D cell culture bridges the gap between in vitro and in vivo studies. These systems

have allowed for significant strides in cell biology through the reestablishment of critical

microenvironmental factors, particularly cell-cell and cell-ECM interactions, in a range of work

including tumor biology, cell adhesion, migration, and epithelial morphogenesis. To improve

data reliability from drug screening and avoid the pursuit of in vivo studies on false-positive

targets, 3D cell culture presents a potentially ideal system to balance cost and reliability.

Unfortunately, hydrogel culture methods remain under-utilized in part because of high reagent

costs and challenges in the manipulation and handling of delicate hydrogel materials.

A number of strategies relying on microchannels have been proposed to address the challenges

of working with 3D cell structures in hydrogels.47

Microfluidics provides the ability to

manipulate sub-microlitre volumes of liquid thereby reducing reagent consumption.

Furthermore, the associated low Reynolds number flow through microfluidic channels allows for

gentle hydrogel handling and reduces subsequent damage to gels during reagent exchange.

18

Microfluidic devices used for 3D cell culture and handling have ranged in complexity, from

systems with integrated valves48

and off-chip pumps49

to passive perfusion systems50,51

exploiting gravity and surface tension forces to drive reagent exchange. While these systems are

undoubtedly useful, the microchannel modality remains challenged for this application by (1)

lack of flexibility in hydrogel geometry and size, (2) channel clogging and limited perfusion

through hydrogels, and (3) tubing dead-volumes.51

Limitations in hydrogel geometry for 3D cell culture in microchannels are inherent to device

design. In continuous flow systems, the channel is completely filled with the hydrogel material,

resulting in gel structures conformal to the channel geometry. In two-phase flow systems the

majority of demonstrations have been restricted to monodisperse spherical or rod-shaped solids

produced by either photo-initiation or thermal cross-linking at a T-junction or by flow-

focusing.52,53

Recent demonstrations by the Doyle group54 have exploited stop-flow lithography

in the high-throughput formation of microgels in a range on geometries. Though these systems

are proving particularly useful in the conception of drug delivery systems, they are limited to UV

initiated cross-linking which can be detrimental to cells and excludes the many temperature

sensitive hydrogel systems (e.g., collagen, Matrigel) that are commonly used in 3D cell culture

studies. Further these systems typically produce hydrogel geometries with length scales of <100

µm (and volumes of < 1 nL), whereas typical 3D cell construct sizes range from 50-1000 µm in

size. DMF liquid handling is an emerging alternative to the paradigm of enclosed

microchannels.14

Recently, in the unique geometries of DMF have been exploited for handling

and addressing of three-dimensional solids such as paper discs for blood screening,55,56

polymer

monoliths for sample extractions,57

and agarose discs for scaffolding applications.58,59

In

19

chapters 4 and 5 of this thesis DMF liquid handling is combined with the virtual microwells

described in chapter 2 developing a novel platform for ‘microgels on-demand’, that allows for

sub-microlitre 3D cell culture.

1.6 Assays and integration

The real benefits of DMF are realized in the automation of multi-step assays and integration of

devices within existing laboratory analytical infrastructure. Live-cell apoptosis assays with cell

seeding followed by stimulation, washing, and staining steps were recently performed on DMF.5

These assays were performed in microliter droplet volumes and did not suffer from cell loss

during reagent exchange, a common problem for such assays when performed in microwell

plates. Multiplexing of these assays on a single device provides for the ability to quickly and

efficiently screen a range of conditions (Figure 1-4C). The use of fluorescent apoptosis markers

allows for direct device integration with a fluorescent plate reader of the type that is common in

research laboratories. In chapter 3 of this thesis, primary cells were cultured for multiple days on

DMF.3 These cells were subsequently stimulated with cytokines and their functional responses

tested in a monocyte adhesion assay. Further, on-device fixing and staining of these cells

followed by epifluorescent microscopy and imaging in high-content screening equipment,

demonstrated device compatibility with microscopy for the acquisition of high-quality images.

A second area where the small volume reagent handling capabilities of DMF is currently being

exploited is mass spectrometry (MS). Upstream sample preparation on DMF has embodied

liquid/liquid extractions for DNA clean up through aqueous immiscible droplets60

and tissue

processing by extraction in isooctane.61

Liquid/solid sample prep strategies on device have

20

included hydrogel immobilized enzymes for proteolytic digestion,59

and solid-phase extraction in

porous polymer monoliths. These methods have proven critical in separating analytes from salts

and contaminants that can cause ion suppression thereby hindering analysis. Further, methods

for direct sample introduction to MS have explored devices patterned on flexible substrates as

foldable emitters62

and embedding glass capillaries between the device top and bottom plates to

facilitate reagent transport into the MS.56

DMF integration with analytical methods remains an

active area of research with other groups coupling these systems with emerging applications

including surface plasmon resonance (SPR)63

and miniaturized tunable droplet lasing systems64

.

1.7 Future of DMF

One of the visions of digital microfluidics for high throughput biological screening is the

eventual development of low-cost ‘smart’ microwell plates to complement or replace automated

liquid handling robotics. Ideally these would function as self-contained cell culture and analysis

units capable of multiplexed cell based assays. This will require the combination of robust low-

cost devices, novel surfaces that are resistant to biofouling, and automation hardware to drive

droplet translation. In the development of this technology groups are finding capabilities of DMF

systems for applications that are not possible utilizing other technologies – including temporal

studies of cellular responses to cytokines and thermally crosslinked microgel formations in

customizable geometries (described in chapters 4 and 5). With an increasing number of research

groups actively pursuing the development of DMF, there remains great potential for this platform

technology to function as novel foundation for HTS and provide essential tools in gaining greater

insights on fundamental biological processes.

21

Chapter 2. Virtual microwells for digital microfluidic reagent

dispensing and cell culture

Summary

Digital microfluidic (DMF) liquid handling includes active (electrostatic) and passive (surface

tension) mechanisms for reagent dispensing. Here we implement a simple and straightforward

Teflon-AF liftoff protocol for patterning hydrophilic sites on a two-plate device for precise

passive dispensing of reagents forming virtual microwells – an analogy to the wells found on a

microtitre plate. We demonstrate here that devices formed using these methods are capable of

reproducible dispensing of volumes ranging from ~80 to ~800 nL, with CVs of 0.7% to 13.8%

CV. We demonstrate that passive dispensing is compatible with DMF operation in both air and

oil, and provides for improved control of dispensed nano- and microlitre volumes when

compared to active electrostatic dispensing. Further, the technique is advantageous for cell

culture and we report the first example of reagent dispensing on a single-plate DMF device. This

method has proven useful for DMF based cell culture and analysis. The technology described in

this chapter presents a platform upon which the remainder of this thesis is built upon.

22

2.1 Introduction

Miniaturization of laboratory procedures for lab-on-a-chip technologies requires on-device

methods that are analogous to standard pipette-based reagent dispensing. This has been realized

in microchannel-based systems through on/off device pumps,65

in-line valves,48

electrokinetic

flow,65

and capillary action,66

providing control of femto- to microlitre volumes. In digital

microfluidics (DMF), a technique in which droplets are manipulated across an array of insulated

electrodes, active14

(electrostatic) and passive67

(surface tension) dispensing modes have been

demonstrated. Here, we introduce an improvement to passive dispensing, with an emphasis on

robustness and reproducibility, and applications in cell culture and analysis.

DMF devices are operated in either single or two-plate geometries. The single plate geometry

typically consists of actuation electrodes with co-planar ground electrodes60

or a suspended

grounded catena.68

In the two-plate format (Figure 2-1A,B), a bottom plate is patterned with

electrodes coated with a dielectric and hydrophobic material, and a top plate comprises a

conductive layer coated with a hydrophobic material. A widely used function of two-plate digital

microfluidics is active dispensing of droplets from reservoirs.14

As shown in Figure 2-1C (panels

i and ii), active dispensing is achieved by actuating a series of electrodes to stretch, neck, and

pinch a droplet off from a reservoir.14

Active dispensing highlights a particularly useful property

of digital microfluidics: reagents and samples can be dispensed reliably and precisely on-

demand. However, as demonstrated in this chapter, inconsistencies arise when dispensing

reagents with varying viscosities.

23

Figure 2-1: Two-plate digital microfluidic (DMF) device design and assembly. (A) Exploded

view of a device, comprising a bottom plate with patterned electrodes and a top plate bearing

patterned hydrophilic sites. (B) Side-view, not to scale. (C) Schematic depicting two reagent-

dispensing mechanisms on DMF. Active dispensing (i & ii) involves electrostatic stretching of a

reagent from the reservoir followed by splitting. Passive dispensing (iii & iv) occurs

spontaneously as a source droplet is translated across the hydrophilic site. The inset is a three-

dimensional depiction of a virtual microwell, VM (i.e., a droplet formed by passive dispensing).

VM volume is dictated by the diameter of the hydrophilic site (d) and the distance between top

and bottom plates (h).

24

An alternative digital microfluidic function called passive dispensing was recently described by

Barbulovic-Nad et al,67

building on similar work by Chen et al.69

Passive dispensing is

implemented using a DMF device surface that is primarily hydrophobic but patterned with

hydrophilic regions. When a source droplet is translated across a hydrophilic site, surface tension

effects result in spontaneous formation of a sub-droplet on the patch (Figure 2-1C, panels iii and

iv). As described previously,41,70–72

passive dispensing is particularly useful for adherent

mammalian cell culture, allowing for cell seeding onto dry hydrophilic sites, as well as for

subsequent media and reagent exchange on droplet-bearing sites. We introduce here a new term

for the cylinder-shaped droplet formed by passive dispensing: a virtual microwell (VM). The

term VM is an analogy to the wells found on a microtitre plate. The "wells" described here are

virtual as they are not confined on the sides like traditional wells, but are defined by the surface

properties of the top and bottom plate. A similar strategy has been described previously73

for

non-microfluidic applications, however this is the first time this concept is being applied within

the context of DMF.

In initial work describing passive dispensing for cell culture,67

hydrophilic patches were formed

by adsorbing extracellular matrix proteins onto Teflon-AF-coated DMF device bottom plate

surfaces. Adaptation of this method for other applications realized several challenges, including:

(1) inconsistent reagent dispensing both initially and during subsequent droplet passages, (2)

protein dissolution and subsequent loss of hydrophilic pad integrity, and (3) difficulty

functionalizing the electrode-bearing surface. Motivated by these challenges, we sought to

develop a simple fabrication protocol for patterning hydrophilic sites directly on device surfaces.

Here we report a new method for forming hydrophilic patches relying on a Teflon lift-off

25

procedure. The method is straightforward and fast, allowing for rapid generation of an array of

individually addressable virtual microwells by passive dispensing. We demonstrate that this

method can be used for two-plate DMF operation in air or oil to (1) reproducibly and precisely

dispense reagents independent of viscosity based solely on device design parameters, (2)

maintain constant droplet volume after subsequent reagent exchanges, and (3) improve cell

seeding when compared with previously published methods. Further we show the first example

of reagent dispensing on a single-plate device. We propose that these new methods will be useful

for a wide range of applications -- particularly those involving adherent cell culture and analysis.

2.2 Methods and Materials

2.2.1 Reagents

Unless stated otherwise, general-use chemicals were from Sigma Aldrich (Oakville, ON,

Canada) or Fisher Scientific Canada (Ottawa, ON, Canada), fluorescent dyes and cell media

components were from Invitrogen/Life Technologies (Burlington, ON, Canada), and

photolithography reagents were from Rohm and Haas (Marlborough, MA). Deionized (DI) water

had a resistivity of 18 MΩ·cm at 25°C.

2.2.2 Two-plate DMF bottom-plate fabrication

Digital microfluidic devices were fabricated in the University of Toronto Emerging

Communications Technology Institute (ECTI) cleanroom facility, using transparent photomasks

printed at 20,000 DPI (Pacific Arts and Designs Inc., Markham, Ontario). Two-plate DMF

device bottom-plates bearing patterned chromium electrodes were formed by photolithography

and etching of commercially available chromium and positive photoresist coated glass slides

(Telic, Valencia, CA). Briefly, substrates were exposed to UV through a mask (8 s, 29.8

26

mW/cm2) and then developed in MF-321 (~2 min). Chromium was etched in CR-4 (~5 min, OM

Group, Cleveland, Ohio), and then substrates were washed with DI water, dried under a stream

of nitrogen. Substrates were then immersed in AZ 300T (3 min) to remove photoresist and then

washed in DI and dried under a stream of nitrogen. This was followed by cleaning in Piranha

solution (10 s, 1:1 conc. sulfuric acid: 30% hydrogen peroxide). Substrates were rinsed in DI

water, then dried under a stream of N2, before dehydrating on a hot plate (165 ºC, 10 min). As

shown in Figure 1A, the bottom-plate design featured an array of 116 actuation electrodes (2.2

2.2 mm ea.) connected to 10 reservoir electrodes (4 4 mm ea.), with inter-electrode gaps of 30-

80 µm. The actuation electrodes were roughly square with 140 µm (peak to peak) sinusoidal

interdigitations. In some experiments, the design also included an array of five 1 mm diameter

optical windows (i.e., circular regions free from chromium) with 9 mm between each window.

Each window straddled the interface between two actuation electrodes. After patterning, the

substrates were immersed for 30 minutes in silanization solution: 3-(Trimethoxysilyl)propyl

methacrylate (Specialty Coating Systems, Indianapolis, IN), diluted to 1% (vol./vol) in 1:1 DI

water:isopropanol. Substrates were air-dried for 30 minutes then washed with isopropanol (IPA)

and dried under a stream of nitrogen. Substrates were then coated with 8 µm of Parylene-C

(Specialty Coating Systems) and 200 nm of Teflon-AF 1600 (DuPont, Wilmington, DE).

Parylene-C was applied using a vapor deposition instrument (Specialty Coating Systems), and

Teflon-AF was spin-coated (1% wt/wt in Fluorinert FC-40, 3000 rpm, 60 s) followed by post-

baking on a hot-plate (165 ºC, 10 min). Each driving electrode and reservoir was connected to a

contact pad on the edge of the substrate. The polymer coatings were removed from contact pads

by gentle scraping with a scalpel to facilitate electrical contact for droplet actuation.

27

For some experiments, hydrophilic sites were formed on ethanol sterilized DMF bottom plates as

reported previously.67

Briefly, 2 µL aliquots of fibronectin (33 µg/mL in DI water) were pipetted

onto the Teflon-AF surface covering optical windows and were allowed to evaporate at room

temperature for ~4 hours. The adsorbed protein spots formed in this manner were roughly

circular with ~1 mm diameter.

2.2.3 Two-plate DMF top-plate fabrication

Two-plate DMF device top-plates were formed from indium tin oxide (ITO) coated glass

substrates (Delta Technologies Ltd, Stillwater, MN). For most experiments, ITO-glass substrates

were coated with Teflon-AF and then were treated with a fluorocarbon lift-off procedure to form

an array of hydrophilic spots (i.e., circular regions of exposed ITO). Briefly, ITO-glass slides

were immersed in RCA solution (6:1:1 DI water: 28% aqueous ammonium hydroxide: 30%

hydrogen peroxide) for 15 minutes at 80°C. After rinsing, drying, and dehydrating, substrates

were spin-coated with Shipley S1811 photoresist (3000 RPM, 60 s) and then post-baked on a hot

plate (2 min, 95°C). The substrates were exposed (10 s, 29.8 mW/cm2) through a mask bearing

an array of five 1.00-, 1.25-, 1.50-, 1.75-, or 2.00-mm-diameter circular features (9 mm between

each feature) and then developed in MF-321. After rinsing and drying, the substrates were flood

exposed (10 s, 29.8 mW/cm2), and then spin-coated with Teflon-AF and post-baked using the

same parameters used for bottom-plate substrates (as above). The substrates were then immersed

in acetone with gentle agitation until the Teflon-AF over the patterned sites was lifted off (5-10

s). After rinsing and drying, the Teflon-AF was reflowed by baking on a hot plate at 165°C,

210°C, and 300°C for 5 minutes at each temperature. For some experiments, unpatterned top-

plates were formed without fluorocarbon liftoff, and were simply spin-coated with Teflon-AF

28

using the same parameters used for bottom-plate substrates (as above).

2.2.4 Two-plate DMF device assembly and operation

Two-plate digital microfluidic devices were assembled with an ITO–glass top plate and a

chromium-glass bottom plate as shown in Figure 2-1B. The two plates were joined by stacking

one, two, or three layers of double-sided tape (each layer ~80 µm), and were aligned such that

the edge of the top plate was adjacent to the outer-edges of the reservoir electrodes on the bottom

plate. In cases in which optical windows and top-plate hydrophilic sites were used, care was

taken to align these features vertically (windows on the bottom plate and hydrophilic sites on the

top plate). Driving potentials, ~300 VRMS for operation in air or ~200 VRMS for operation in oil,

were generated by amplifying the sine wave output of a function generator (Agilent

Technologies, Santa Clara, CA) operating at 18 kHz. Each reagent was loaded onto the device by

pipetting an aliquot onto the bottom plate at the edge of the top plate, and simultaneously

applying driving potential to the closest reservoir electrode (relative to the ITO electrode on the

top plate) to draw the fluid into the reservoir. Thereafter, droplets were actively dispensed,

moved, and merged by applying driving potentials to sequential actuation electrodes on the

bottom plate relative to the ITO electrode on the top plate as described previously.14

In all DMF

experiments, reagent solutions were supplemented with 0.02% Pluronics F68.74

2.2.5 Single-plate DMF device fabrication, assembly, and operation

The one-plate DMF device design consisted of twelve 3 x 3 mm square electrodes adjacent to a

linear 1 mm wide ground electrode with 40 µm between each electrode. Devices were coated

with Parylene-C using the same method described for bottom plates of two-plate devices (as

29

above). Devices were then coated with Teflon-AF bearing 1 to 2 mm diameter circular

hydrophilic sites using a modified liftoff procedure. Briefly, 10 nm of chromium was deposited

onto the Parylene by electron beam evaporation, which was then patterned into circular sites by

photolithography and etching using parameters described for bottom plates of two-plate devices

(as above). Devices were spin coated with Teflon-AF 1600 (1% wt/v in FC-40, 3000 RPM, 1

min), baked for 10 min at 165°C then flood exposed (10 s, 29.8 mW/cm2). Devices were

immersed in acetone with gentle agitation until Teflon-AF lifted off (~5-10 min) revealing a

pattern of circular chromium features. Devices were rinsed with DI, dried under a stream of

nitrogen, then baked on a hot plate for 10 min at 165°C. Single-plate devices were loaded by

pipetting 20 µL of reagent directly onto the outermost driving electrode. Sine wave driving

potentials of ~600 VRMS at 18 kHz were applied with the same amplified function generator

described above. Droplets were made to translate across the device by potentiating sequential

square electrodes relative to the linear electrode (held at ground) as described previously.60

2.2.6 DMF dispensing experiments

Devices bearing droplets were imaged with a CCD camera (Basler, Ahrensburg, Germany)

mounted above the device. For two-plate devices, ImageJ software was used to estimate the

apparent cross-sectional area of each droplet (typically circular but in some cases in the shape of

an irregular polygon) and volume (knowing the intra-plate spacer thickness). For one-plate

devices, each device was weighed on a microbalance before and after dispensing (after removal

of the remainder of the source droplet with a tissue), allowing for estimation of dispensed

volume on the basis of mass. The reagents evaluated included phosphate buffered saline (PBS)

with 0.2% blue food dye, 0-65 wt% sucrose solutions prepared in DI water (with viscosities from

30

literature values75

), and Dulbecco’s Modified Eagle Medium (DMEM) with 10% fetal bovine

serum (FBS). At least three replicates were performed for all conditions.

In each experiment, an aliquot of the appropriate reagent was loaded into a device, and one of a

number of conditions was evaluated. (1) Active dispensing on a two-plate device. A unit droplet

(i.e., a droplet covering one actuation electrode) was actively dispensed onto actuation electrodes

and the volume was estimated. (2) Passive dispensing onto top-plate hydrophilic sites in air on a

two-plate device. A unit droplet was actively dispensed as in (1) above and then translated over a

hydrophilic site, and the volume of the passively dispensed droplet was estimated. In some

instances, two unit droplets were actively dispensed and merged, then the combined droplet was

translated over the hydrophilic site, and the volume of the passively dispensed droplet volume

was estimated. For all spot sizes, dry dispensing (i.e., passive dispensing onto sites not bearing a

droplet), and wet dispensing (i.e., passive dispensing onto sites bearing a droplet from a previous

dispensing experiment) were evaluated. (3) Passive dispensing onto top-plate hydrophilic sites in

oil on a two-plate device. Droplets were first passively dispensed in air (dry dispensing) as in (2),

above. The entire device (i.e., all of the space between the top and bottom plates not occupied by

a droplet) was then filled with light mineral oil. Wet dispensing was then evaluated as in (2),

above. (4) Passive dispensing onto bottom-plate hydrophilic sites in air on a two-plate device. A

unit droplet was actively dispensed as in (1) above, and then translated over the hydrophilic site

(i.e., a patch of adsorbed protein on the bottom plate), and the volume of the passively dispensed

droplet volume was estimated. (5) Passive dispensing on a one-plate device. A 20 µL droplet

was translated across a patterned hydrophilic site. A sub-droplet was generated by passive

dispensing (after which, the volume was estimated), as the main droplet was actuated away.

31

2.2.7 Cell Culture and experiments

Marbin Darby canine kidney (MDCK) epithelial cells were kindly provided by Dr. N. Tufenkji

(McGill University). MDCKs were cultured in DMEM supplemented with 10% FBS, 100 U/mL

penicillin, 100 ug/mL streptomycin. Cells were incubated at 37°C in a humidified incubator

containing 5% CO2. 5 µL aliquots of cell suspensions (1 106 cells/mL) in media were pipetted

onto reservoir electrodes of two-plate devices bearing optical windows on the bottom plate. Unit

droplets were actively dispensed onto the electrode array, and translated across hydrophilic sites,

(formed either by liftoff on the top-plate or fibronectin absorption on the bottom-plate), resulting

in passive dispensing. Devices were then incubated at 37°C in a humidified incubator containing

5% CO2 for 18 hours. Images of cells were acquired through the optical windows by light

microscopy and the numbers of cells dispensed were enumerated using ImageJ.

2.3 Results and Discussion

2.3.1 Lift-off patterning

DMF devices are typically coated with fluorocarbon (FC) polymers such as Teflon-AF (DuPont),

CYTOP (Asahi), or Fluorad (3M). These materials have desirable properties including low

surface energy, broad chemical resistance, thermal stability, and biocompatibility. Early DMF

applications were implemented in devices bearing homogenous FC surfaces for applications

including suspension cell culture,40

PCR,76

enzymatic assays,77

and DNA sequencing.78

More

recently, DMF devices have been combined with heterogeneous surfaces (bearing different

chemical functionalities) for more sophisticated applications. These methods can be sub-divided

into those relying on modifications of the device surface itself67

or by incorporation of external

materials with heterogeneous surface properties such as magnetic beads79

or polymer plugs.57

In

32

this paper, we focus on the former -- heterogeneous patterned device surfaces. As described in

the introduction (and depicted in Figure 2-1C), DMF devices with surface modifications are

particularly useful for a form of fluidic manipulation called passive dispensing.40

We report here a technique to form DMF devices that are globally coated with Teflon-AF, but

periodically patterned with hydrophilic spots. In most of the work reported here, the hydrophilic

spots were formed from exposed indium tin oxide (ITO) on the top plate of a DMF device as

illustrated in Figure 2-1B. Our new technique is similar to that described by Chen et al. and

Malic et al.80

for forming patterned Teflon-AF on ITO and gold surfaces, respectively.

Significant trial and error was required to develop techniques that were reproducible with

particularly important results being inclusion of an RCA cleaning step for improved adhesion of

Teflon-AF to ITO and an extra UV exposure step to assist in photoresist removal. We

characterized these surfaces by x-ray photoelectron spectroscopy finding similar indium, tin, and

oxygen compositions on the liftoff sites as compared to untreated ITO (Figure 2-2 and Table

1-1). These measures and others (described in detail in the experimental section) form a robust

and repeatable method that we have now used to pattern hundreds of substrates bearing circular

structures with near-perfect pattern fidelity.

2.3.2 Passive dispensing and virtual microwells

The new methods for patterning surfaces described above were developed to facilitate robust

formation of virtual microwells by passive dispensing. As shown in Figure 2-3A, the simplest

form of passive dispensing can be called "dry" passive dispensing, in which a VM is formed on

an empty hydrophilic site. We evaluated the effects of varying gap spacing between

33

Figure 2-2: X-ray photoelectron spectroscopy evaluation of patterned surfaces. To evaluate the

chemical composition of exposed hydrophilic sites and adjacent Teflon surfaces XPS

measurements were taken on patterned and unpatterned surfaces for comparison: (A) ITO

composition at hydrophilic sites, (B) Teflon on patterned slides, (C) unprocessed ITO surfaces,

and (D) unprocess Teflon surfaces.

Table 2-1: Elemental identification and quantification of species detected on patterned DMF

surfaces using XPS. Samples labels refer to Figure 2-2. Sample (Atomic %)

Species A B C D

C 40.73 38.30 36.30 31.00

F 0.31 50.26 52.38 0.00

N 0.68 0.11 0.01 0.10

S 1.26 0.01 0.00 0.00

Sn 2.61 0.00 0.00 4.14

In 17.22 0.00 0.00 26.77

O 35.83 11.12 11.29 37.86

Si 0.34 0.03 0.01 0.13

Ca 1.02 0.16 0.01 0.00

34

the top and bottom plates and the diameter of the hydrophilic site on dry passive dispensing. As

shown in Figure 2B, by varying the hydrophilic site diameter from 1 mm to 2 mm and the inter-

plate gap height from 80 µm to 240 µm, VMs with volumes ranging from ~80 nL to ~800 nL

were formed. The precision of these volumes varied within the range of 0.7% to 13.8% for all

conditions tested. The CVs increased with greater dispensed volume (either higher gap spacing

or larger hydrophilic surface area).

In initial experiments, we observed that a single actively dispensed droplet (i.e., Figure 2-1C,

frames i-ii) was not always sufficiently large to serve as the source droplet for dry passive

dispensing. In such cases, the VM did not properly separate from the source droplet, or the

remainder of the source droplet (after forming the VM) was too small to actuate away. Thus, for

the conditions in Figure 2-3B labeled with an asterisk (*), two droplets were actively dispensed

and subsequently merged, and this combined volume served as the source droplet for passive

dispensing. This observation led us to develop a quantitative criterion predictive of passive

dispensing success, which we call the "virtual microwell number," Nvm, which is defined in terms

of the area of the square actuation electrodes (Ae), the area of the circular hydrophilic site (Ahs),

and the distance between top and bottom plates (h):

35

Figure 2-3: Dry passive dispensing to form virtual microwells. (A) Video stills (top-to-bottom)

depicting dry passive dispensing. The dashed circle in panel (i) indicates the position of the

hydrophilic site. (B) Volumes of droplets dispensed in dry passive dispensing as a function of

spacer height and hydrophilic site diameter (n = 5). Asterisks (*) indicate that source droplets were

formed from two actively dispensed droplets. Error bars are 1 S.D. (C) Parameter NVM calculated

for each experimental condition in (B). The shaded region, NVM < 2, indicates conditions in which

two actively dispensed droplets were required to generate the source droplet for successful passive

dispensing.

36

hA

AN

hs

evm Eqn. 2.1

Figure 2-3C summarizes Nvm across a range of device and feature parameters. We observe for all

experiments where Nvm > 2, a single unit droplet actively dispensed from the reservoir was

sufficient for successful generation of the VM. For Nvm < 2, a single unit droplet was insufficient

for successful passive dispensing. This is mostly consistent with the observations described by

Chen et al.,69

with a discrepancy observed for cases when Nvm is close to 2. We propose that this

discrepancy may be attributed to differences in device design and operation. Regardless, we

anticipate that Nvm will be a useful heuristic in the design of VMs on DMF devices in the future.

For the majority of applications it is of interest to exchange reagents in VMs. We term this type

of exchange "wet" passive dispensing, which is implemented when a source droplet is actively

dispensed and then translated across a previously formed VM, displacing its original contents

(Figure 2-4A). Barbulovic-Nad et al.67

demonstrated that after three such exchanges, 100% of the

content of the original VM is replaced. We evaluated VMs for hydrophilic sites with diameters

ranging from 1000 to 2000 µm for the ability to repeatedly dispense volumes to sites bearing

VMs. As shown in the gray bars in Figure 2-4B, the precision in wet dispensing is very high for

small sites, with CVs of 1.8%, 1.5%, and 0.7% for 1000 µm (126 nL), 1250 µm (196 nL), and

1500 µm (283 nL) diameter hydrophilic sites, respectively. Larger hydrophilic sites were

associated with lower precision, with CVs of 12% and 7% for 1750 µm (385 nL) and 2000 µm

(502 nL) diameter hydrophilic sites, respectively. Regardless, these data indicate that given

37

Figure 2-4: Wet passive dispensing to exchange fluid in a virtual microwell. (A) Video stills

(top-to-bottom) depicting wet passive dispensing in which the virtual microwell contained blue

dye at the hydrophilic site (i) and a red dye source droplet is actuated across the virtual

microwell displacing the blue droplet (ii-v). (B) Multiple passes of reagent across virtual

microwells with varying diameters for 160 µm spacer height. The gray and white bars represent

devices operated with a surrounding matrix of air and mineral oil, respectively. Error bars are 1

S.D.

38

volumes can be repeatedly dispensed to a given site multiple times with good (and for low

volumes, excellent) precision.

The data above (and in most of the experiments described here) were generated using devices in

which droplets were surrounded by a matrix of air. An alternative format is to fill devices such

that droplets are surrounded by a matrix of oil, which has the benefit of lower voltage

requirements, reduced surface fouling, and decreased droplet evaporation.80

We demonstrate

here the implementation of passive dispensing in oil-filled DMF devices. Interestingly, we found

dry passive dispensing in oil to be impossible. We speculate that this is because a thin film of oil

film forms over the hydrophilic site and prevents hydrophilic interactions with aqueous droplets.

In contrast, we found that wet passive dispensing in oil was straightforward when VMs were first

loaded in air by dry passive dispensing and the devices were then filled with light mineral oil

(which did not displace the aqueous droplets from the VMs) for wet dispensing, as shown in the

white bars in Figure 2-4B. The VMs formed in oil were in 1.1-fold to 1.25-fold larger the initial

dry dispensed volume in air. We attribute this phenomenon to the increased viscous forces

between the droplet and the filler medium.22

With multiple passes, the oil-associated volume was

maintained consistently, with CVs ranging from 2.8% to 9.4% for volumes of 240 nL to 810 nL.

We propose that the capacity to combine passive dispensing with oil-filled devices will be useful

for a range of different applications benefitting from the use of oil to facilitate droplet motion

and reduce the effects of potential evaporation.

The data above (and in most of the experiments described here) were generated using two-plate

DMF devices in which droplets are sandwiched between a top and bottom plate (Figure 2-1). In

39

Figure 2-5: Single-plate DMF passive dispensing. (A) Picture of a single-plate device depicting

a source droplet and a passively dispensed droplet. (B) Schematic depicting the single-plate

device geometry.

Figure 2-6: Active and passive dispensing as a function of reagent viscosity. Sucrose solutions

of varying viscosity were dispensed on DMF either by active or passive dispensing onto 1500

µm diameter hydrophilic sites (n = 6). Dispensed volumes are plotted as a function of solution

viscosity. Error bars are ± 1 S.D., 95% confidence intervals are indicated by shaded regions, and

the mean dispensed volume for each dispensing mechanism is indicated by a solid horizontal

line.

40

the alternative single-plate device format, the larger droplet volume-to-electrode-area ratio

results in lower actuation forces relative to two-plate DMF; thus as far as we are aware, there

have been no reports of reagent dispensing (of any kind) on single-plate DMF devices. Here, we

report the extension of the concept of passive dispensing on hydrophilic sites to single-plate

DMF devices (Figure 2-5). Applying fluorocarbon lift-off to single-plate DMF devices provides

the ability to dispense reagents in this format. The hydrophilic sites in such systems were formed

by evaporation of chromium (and subsequent patterning) on the top of the Teflon surface of a

complete single-plate device. For hydrophilic sites with diameters of 1250 µm and 1500 µm, the

droplets dispensed from 20 µL source droplets had volumes of 330 nL ± 35 nL and 420 nL ± 55

nL (CVs of 11 and 13%, respectively).

2.3.3 Active vs. passive dispensing

Active dispensing is regarded as the standard technique for reagent dispensing on DMF. In active

dispensing, the liquid is stretched from a reservoir by electrostatic manipulation and then necked

prior to splitting. Fouillet et al.81

reported a CV of below 4% for active droplet dispensing of

reagent into oil-filled DMF devices, similar to pipettes, where precision is reported at under 6%

CV for 0.1 to 2.5 uL. Actively dispensed volumes are limited by device geometry; different

electrode sizes are required to achieve dispensing of different volumes.57

Further, when we

examined the active dispensing of a range of sucrose solutions with varying viscosity from 0 cP

to ~150 cP, we found poor repeatability, with errors of up to 30% and a 95% confidence interval

across all viscosities of 0.2 mm3 (red circles in Figure 2-6). In comparison, VM volumes formed

by passive dispensing were relatively independent of viscosity, with calculated 95% confidence

41

intervals for dispensed volume of 0.02 mm

3. In the future, we propose that the integration of

different sizes of hydrophilic sites on device will make it straightforward to access a broad range

of reagent volumes, and improve precision and accuracy of reagent dispensing independent of

viscosity.

2.3.4 Lift-off vs. protein absorption for passive dispensing

The liftoff-based techniques for forming hydrophilic patches on device top plates described here

were developed as a result of our dissatisfaction with methods relying on hydrophilic patches

formed from adsorbed proteins,67

for the reasons listed in the introduction. Here, we report a

comparison of the two systems for the ability to passively dispense droplets containing

suspended cells, and the ability of the cells to spread on the device surface. After performing

independent trials of dispensing MDCK cells suspended in cell culture media, we found lower

cell numbers and greater variability in the case of sites formed by protein spotting (cell number

mean = 24 cells, cell number CV = 79%) as compared to those formed by fluorocarbon liftoff

(cell number mean = 70 cells, cell number CV = 19%) (Figure 2-7A). Furthermore, as shown in

Figure 2-7B, there were no significant differences in cell morphology for the two types of

systems, which supports literature reports of ITO as being a suitable surface for cell culture.82

In addition to improved reproducibility in dispensing, the new technique reported here has

additional benefits for cell culture in DMF, including increased electrical isolation of cells from

the actuation electrodes and compatibility with long-term culture. In DMF systems such as those

reported here, the majority of the voltage drop occurs across the dielectric coating of the bottom-

plate;40

however, localized charge densities at the device surface may vary, which might result in

42

Figure 2-7: Comparison of hydrophilic sites formed by adsorbed protein (on the bottom plate)

vs. liftoff (on the top plate) for dispensing cells into virtual microwells. (A) Results of five trials

seeding MDCK cells (5 105 cells/mL) by passive dispensing. (B) Bright-field images of

MDCK cells seeded on fibronectin coated Teflon and indium tin oxide after 6 hours. Scale bar =

50 µm.

43

augmented transcriptional profiles within cultured cells. Further, localized heating in the

dielectric layer might result in potentially deleterious cellular effects. Decoupling the cell culture

site (by moving it to the top plate) from the electrode-bearing substrate (on the bottom plate) may

dampen these effects. In the case of long-term culture, dielectric coatings are prone to failure due

to accumulation of charge and moisture infiltration during incubation. Cell culture on the top-

plate allows for the replacement of defective bottom-plates (with cells grown continuously on

top-plates) without compromising the experiment being performed. We propose that this

arrangement will be useful for the long-term culture of sensitive cell types, particularly stem and

primary cells.

2.4 Conclusion

We present the utility of Teflon liftoff for improving digital microfluidic functionality. This

precise method for patterning hydrophilic sites on hydrophobic DMF device surfaces resulted in

multiple advances, including: (1) formation of virtual microwells for precise reagent dispensing,

(2) passive dispensing in air and oil filled devices, (3) the first demonstration of passive

dispensing on a single-plate device, and (4) improved surfaces for cell culture and other

heterogeneous assays. We anticipate this new method will be useful for DMF-based techniques

applied to a broad range of applications.

44

Chapter 3. A digital microfluidic platform for primary cell

culture and analysis

Summary

Digital microfluidics (DMF) is a technology that facilitates electrostatic manipulation of discrete

nano- and microlitre droplets across an array of electrodes, which provides the advantages of

single sample addressability, automation, and parallelization. There has been considerable

interest in recent years in using DMF for cell culture and analysis, but previous studies have used

immortalized cell lines. We report here the first digital microfluidic method for primary cell

culture and analysis. A new mode of “upside-down” cell culture was implemented in by

patterning the top plate of a device using a fluorocarbon liftoff technique. This method was

useful for culturing three different primary cell types for up to one week, as well as

implementing a fixation, permeabilization, and staining procedure for F-actin and nuclei. A

multistep assay for monocyte adhesion to endothelial cells (ECs) was performed to evaluate

functionality in DMF-cultured primary cells and to demonstrate co-culture using a DMF

platform. Monocytes were observed to adhere in significantly greater numbers to ECs exposed to

tumor necrosis factor (TNF)-α than those that were not, confirming that ECs cultured in this

format maintain in-vivo-like properties. The ability to manipulate, maintain, and assay primary

cells, demonstrates a useful application for DMF in studies involving precious samples of cells

from small animals or human patients.

45

3.1 Introduction

There are two types of mammalian cells that are commonly used in biomedical research:

immortalized cell lines and primary cells. Immortalized cell lines can be grown in vitro for many

generations, spanning many months-to-years. These cells are straight-forward to grow and

maintain, but often have phenotypes that differ significantly from those of cells in vivo. In

contrast, primary cells are used immediately after isolation from animal tissue, and therefore are

much closer to in vivo phenotype. Unfortunately, primary cells have several limitations for

regular use in the laboratory. In long-term studies involving animal models of disease, primary

cells are typically available only in limited quantities (e.g., with monthly or yearly isolations).

The process of primary cell isolation can be laborious and costly, requiring expensive reagents

and hours-to-days of work depending on the cell type. Furthermore, due to their limited number

of population doublings, primary cells can only be used for a short period of time in the

laboratory. These factors make primary cells an attractive target for miniaturized tools to reduce

costs and for automated cell culture and analysis.

Microfluidic channels are the most popular technology used for miniaturization. Primary cell

culture in microfluidic channels has been demonstrated repeatedly with applications including

cell migration,1-3

adhesion,4-6

shear stress,7-9

cell sorting,10

and cell-based screening assays.11

However, microchannel-based systems often require pumps or other external apparatus (with

noted exceptions12

) for applications involving cells. This increases reagent/sample consumption,

as such systems require macro-scale tubing and interconnects, which inherently contributes

unwanted dead volumes. An additional problem associated with interconnects and other world-

to-chip interfaces is the presence of bubbles, which can disturb the local fluid flow within

46

microchannels and can damage cells as a result of the high interfacial energy at the gas-liquid

interface. Removing bubbles can be difficult, requiring complex degassing mechanisms or

bubble traps.13

Digital microfluidics (DMF) is an alternative platform to conventional enclosed microchannels

that is capable of manipulating discrete liquid droplets on an array of patterned electrodes.14

In

DMF, droplets can be controlled individually or in parallel to provide precise spatial and

temporal control of reagents. Typical volumes for droplets can range from nanolitres to

microlitres, and because there is no dead volume, these systems are well suited for minimal

reagent/sample consumption. Moreover, because there are no open reservoirs or tubes and

interconnects, devices can be readily flipped, allowing for convenient use of both sides of each

device for imaging. Finally, unlike enclosed microchannels, in non-oil-filled DMF systems,

bubble nucleation and growth are non-existent. Previous studies15-21

have demonstrated that

mammalian cells can be cultured and/or analyzed on DMF platforms, but all of the previous

work used immortalized cell lines.

Here, we report the first application of DMF to the culture and analysis of primary cells. Three

phenotypically different cell types isolated from pig blood vessels (aortic endothelial cells) and

heart valves (aortic valve endothelial cells and aortic valve interstitial cells) were cultured and

analyzed on a DMF platform. The devices and methods reported here use a new mode of

"upside-down" culture in virtual microwells22

formed by a patterned DMF top plate. Cells were

cultured on multiple sites per device for up to one week. With minimal reagent use, primary

mammalian cells were fixed, permeabilized and stained on a DMF device. Furthermore, a co-

47

culture system for growing an analyzing endothelial cells and monocytes was developed; this is

the first co-culture system that we are aware of in DMF. The co-culture system was used to

implement a monocyte adhesion assay, which confirmed that intricate signaling mechanisms

were retained by primary cells cultured on this new digital microfluidic platform.

3.2 Methods and Materials

3.2.1 Reagents and Materials

Unless stated otherwise, materials were purchased from Fisher Scientific Canada (Ottawa, ON,

Canada). General-use chemicals were from Sigma Aldrich (Oakville, ON, Canada), fluorescent

dyes were from Invitrogen/Life Technologies (Burlington, ON, Canada), and photolithography

reagents were from Rohm and Haas (Marlborough, MA). Deionized (DI) water had a resistivity

of 18 MΩ·cm at 25°C.

3.2.2 DMF Device Fabrication and Operation

Digital microfluidic devices were fabricated in the University of Toronto Emerging

Communications Technology Institute (ECTI) cleanroom facility, using transparent photomasks

printed at 20,000 DPI (Pacific Arts and Designs Inc., Markham, Ontario). Glass DMF device

bottom-plates bearing patterned chromium electrodes were formed by photolithography and

etching as described previously.15

As shown in Figure 3-1, the design featured an array of 116

actuation electrodes (2.2 mm x 2.2 mm ea.) connected to 10 reservoir electrodes (4 mm x 4 mm

ea.), with inter-electrode gaps of 30-80 µm. The actuation electrodes were roughly square with

interdigitated borders (140 µm peak to peak sinusoids). The design also included an array of five

1 mm diameter optical windows (i.e., circular regions free from chromium) with 9 mm between

48

Figure 3-1: (A) Photograph of DMF device designed for primary cell culture and analysis. A

series of droplets (coloured with red dye for visualization) are positioned at patterned hydrophilic

sites on a device. (B) Schematic of device geometry. The top plate is patterned by a liftoff

procedure to expose hydrophilic sites. The bottom plate bears an array of individually

addressable electrodes with patterned optical windows for imaging. (C) Top and side view

schematic of passive dispensing on hydrophilic sites. (i-ii) A droplet is manipulated to the

hydrophilic site. By actuation of subsequent electrodes the droplet is (iii) stretched then (iv)

passively dispensed, forming a virtual microwell. (D) Side view schematic of device orientation

during experimentation. Devices are maintained right-side up during droplet actuation and are

positioned upside-down during all incubations.

49

each window. As illustrated in Figure 3-1C, each window straddled two actuation electrodes.

After patterning the electrodes, the substrates were coated with 7 µm of Parylene-C (Specialty

Coating Systems, Indianapolis, IN) and 200 nm of Teflon-AF (DuPont, Wilmington, DE).

Parylene-C was applied using a vapor deposition instrument (Specialty Coating Systems), and

Teflon-AF was spin-coated (1% wt/wt in Fluorinert FC-40, 3000 rpm, 60 s) followed by post-

baking on a hot-plate (165 ºC, 10 min). The polymer coatings were removed from contact pads

by gentle scraping with a scalpel to facilitate electrical contact for droplet actuation.

DMF device top-plates were formed from indium tin oxide (ITO) coated glass substrates (Delta

Technologies Ltd, Stillwater, MN) that were coated with Teflon-AF (200 nm, as above). A lift-

off process was used to form an array of 1.5 mm diameter openings of exposed ITO (9 mm

between each opening) through the Teflon-AF using methods developed for this purpose.22

Digital microfluidic devices were assembled with an ITO–glass top plate and a chromium-glass

bottom plate. Prior to assembly, the two plates were sterilized by immersing in 70% ethanol (10

min) and then air dried. The hydrophilic sites (exposed ITO) on the top plate were aligned

visually to the optical windows on the bottom plate, and the two plates were joined by a spacer

formed from four pieces of double-sided tape (total space between plates ~280 µm). Driving

potentials (~280 VRMS) were generated by amplifying the sine wave output of a function

generator (Agilent Technologies, Santa Clara, CA) operating at 18 kHz and were applied

between the top plate (ground) and sequential electrodes on the bottom plate via the exposed

contact pads. Pluronics F68 (0.02% wt/vol) was added to all reagents used with digital

microfluidics (excluding solutions of Triton X-100) to facilitate droplet movement.23

A

50

In addition to the standard digital microfluidic operations24

(i.e., active droplet translation, active

droplet dispensing from reservoirs, etc.), the devices supported a phenomenon known as passive

dispensing.15

As illustrated in Figure 3-1C, in passive dispensing, a source droplet is translated

across a hydrophilic site, and surface tension effects result in spontaneous formation of a sub-

droplet. In the devices with the dimensions described here, source droplets were 1.4 µL and

passively dispensed droplets were 0.5 µL; as reported elsewhere,22

the volumetric reproducibility

for passive dispensing for these dimensions is excellent, with a CV of ~1.2%. As described

below, passive dispensing was used for all DMF operations for primary cell culture and analysis.

3.2.3 Primary Cell Isolation and Maintenance

Porcine aortic endothelial cells (PAECs) isolated from pig thoracic aortas were kindly donated

from Lowell Langille (University of Toronto).25

Porcine aortic valve endothelial cells (PAVECs)

and porcine aortic valvular interstitial cells (PAVICs) were isolated as described previously.26, 27

PAECs were cultured in M199 (Wisent, St. Bruno, QC, Canada) supplemented with 5% cosmic

calf serum (Fisher Scientific Canada), 5% fetal bovine serum (FBS) (Fisher Scientific Canada),

and 1% penicillin-streptomycin (P-S) (Sigma Aldrich). PAVECs and PAVICs were cultured in

M199 and Dulbecco’s modified eagle’s medium (DMEM) (Wisent), respectively, each

supplemented with 10% FBS and 1% P-S. Cells were cultured in T75 flasks until 80% confluent,

then trypsinized, centrifuged, and resuspended at approximately 105-10

6 cells/mL in the

appropriate completed culture medium (with M199 or DMEM, as above) to form a cell

suspension for use with DMF.

51

3.2.4 DMF Cell Culture

Five 5 µL aliquots of cell suspensions were pipetted onto the reservoir electrodes, and then five

1.4 µL droplets (one per reservoir) were actively dispensed by applying potentials to a series of

actuation electrodes adjacent to each reservoir. These 1.4 µL cell-containing droplets were

driven to the hydrophilic spots patterned on the top plate such that 0.5 µL droplets were

generated by passive dispensing (Figure 3-1C). The devices were then inverted (with the top

plate on the bottom) (Figure 3-1D) and were placed in a homemade humidified chamber (a Petri

dish containing dampened Kimwipes to prevent evaporation) in an incubator at 37 oC and 5%

CO2 for 12 h. This “incubation state” (i.e., top plate on the bottom in a humidified chamber in a

cell culture incubator) was used for all incubation steps described herein. Periodically, devices

were removed from the incubator, flipped to orient each device with the ITO top-plate on the top

(such that the device was upright) and used for droplet movement. Afterwards, devices were

returned to the incubation state. For cell culture, new droplets of media were delivered to cells

every 12-16 hours until cells were ~70-80% confluent.

3.2.5 DMF Staining and Microscopy

For imaging without staining, primary cells cultured on DMF were imaged using an inverted

CKX41 microscope (Olympus, Markham, ON, Canada) in phase-contrast mode. For comparison,

cells were also cultured on tissue culture treated polystyrene (TCPS) flasks and imaged. For

imaging of stained cells, after ~70-80% confluence was reached on DMF devices, primary cells

were washed by dispensing at least two 1.4 µL droplets of phosphate buffered saline (PBS)

across the virtual microwell sites (displacing the existing droplets with fresh 0.5 µL volumes).

Cells were fixed and permeabilized by dispensing and actuating three 1.4 µL droplets across the

52

cells (in series) of (a) 10% (v/v in DI water) neutral buffered formalin (NBF) for 5 minutes, (b)

PBS, and (c) 0.01% (v/v in PBS) Triton X-100 for 5 minutes. The cells were then washed (two

droplets of PBS as above), and 1.4 µL droplets containing FITC-labeled phalloidin (0.1 mg/mL

in PBS) were actively dispensed from reservoirs and actuated across the cell culture site such

that 0.5 µL sub-droplets were passively dispensed and then incubated for 45 minutes at room

temperature. The cells were then washed in PBS (as above), and 1.4µL droplets containing

Hoechst (1 µg/mL in PBS) were driven across the c -droplets

were passively dispensed and then incubated for 5 minutes at room temperature, and then

washed again with PBS (as above). Cells on DMF devices were imaged by flipping them (such

that the top plate was on the bottom) using an IX-71 microscope (Olympus) in fluorescence

mode.

3.2.6 DMF Monocyte Adherence Assay

THP-1 monocytes (ATCC, Manassas, WA) were cultured in suspension off-chip in RPMI 1640

medium (Invitrogen/Life Technologies) completed with 10% FBS and 1% P-S. Prior to

experiments, monocytes were centrifuged, resuspended in media containing Hoechst (0.2 µg/ml

in complete medium), incubated for 30 minutes, and then centrifuged and resuspended in fresh

complete medium at 106

cells/mL. PAECs grown to confluence on DMF devices were incubated

with passively dispensed 0.5 µL droplets containing 0 or 25 ng/mL tumour necrosis factor alpha

(TNF)-α (Invitrogen/Life Technologies) in complete medium for 4 hours in the incubation state

(see above). Cells were then rinsed by passively dispensing two 0.5 µL droplets of PBS,

followed by passive dispensing of one 0.5 µL droplet containing calcein AM (2 µM in PBS

containing Ca2+

and Mg2+

) and storing for 15 minutes in the incubation state. Cells were then

53

rinsed by passively dispensing two 0.5 µL droplets of PBS, followed by passive dispensing of

one 0.5 µL droplet of complete culture medium and incubating for 30 min in the incubation state.

0.5 µL droplets containing Hoechst-labeled monocytes were then delivered to the PAECs by

passive dispensing and stored for 10 min in the incubation state. Two droplets of PBS were used

to wash the cells (as above), and the cells were then evaluated using an IX-71 microscope for

monocyte adhesion. One central image per hydrophilic spot was collected and images were

analyzed for monocyte number. Briefly, IMAGEJ software was used to convert images to binary

and the “analyze particles” function was used to count the cells.


3.3.1 Digital Microfluidic Primary Cell Culture

We present here the first digital microfluidic platform capable of culturing and analyzing

primary cells, shown in Figure 3-1. PAECs, PAVECs, and PAVICs were chosen as model cell

types because of their importance in cardiovascular biology.4, 26-29

Although these cell types are

found in close proximity anatomically, they represent three distinctly different phenotypes.30

Moreover, PAVECs are an especially interesting target because they are challenging to isolate

and culture in vitro; under improper culture conditions, they display altered morphologies,

function and short-term viability.26, 29, 31

We hypothesize that if DMF is useful for culturing,

handling, and analyzing these different types of cells (particularly, the sensitive PAVECs),

similar methods may be applicable to cells derived from a wide range of tissue types.

PAECs, PAVECs, and PAVICs are adherent cells -- that is, they attach, spread, and grow on

solid surfaces. There have been three previous reports15, 20, 21

of culture of adherent cells on DMF

54

platforms. As listed in Table 1, the new methods reported here share a number of similarities and

differences with those reported previously. The most notable similarity is that each of these

systems is capable of supporting a phenomenon known as passive dispensing. Passive dispensing

is represented in Figure 3-1C; when an aqueous droplet is driven across a hydrophilic site, a

smaller droplet, which we call a "virtual microwell,"22

is spontaneously formed and left behind.

Passive dispensing to form virtual microwells is a unique feature of digital microfluidics, and

serves as a convenient mechanism to seed, culture, and analyze adherent cells.

The most important difference between the current system and those reported previously15, 20, 21

is the new device format and orientation. The methods reported here rely on hydrophilic sites

formed on the device top plate, which led us to implement a new method of "upside-down" cell

culture in virtual microwells (Figure 3-1D). In this scheme, devices are stored for most of the

time upside-down (i.e., top plate on the bottom) which allows the cells to adhere, spread, and

proliferate. At designated periods, devices are flipped to standard configuration (i.e., ITO plate

on the top) for droplet manipulation, but after experiments, the devices are returned to the

inverted state. This arrangement is advantageous for a number of reasons. First, it allows for cell

growth on hydrophilic sites formed from regions of exposed ITO22

rather than the adsorbed

proteins15

or peptides20, 21

used previously. In initial experiments with primary cells grown on

adsorbed fibronectin on DMF device substrates, we observed that the cells had unexpected

morphologies, whereas on ITO surfaces, cells had morphologies that are similar to those grown

on conventional TCPS substrates. Second, this device arrangement de-couples the active portion

of the digital microfluidic device (i.e., the insulating layer on the bottom plate which allows for

the buildup of charge necessary for droplet movement32

) from the cells. The insulating layers on

DMF devices are prone to failure over time because of dielectric breakdown, and the upside-

55

down culture arrangement allows for the possibility of replacing a used/defective bottom plate

with a fresh one between experiments (note that this putative feature was not used in any

experiments reported here). We propose that this arrangement will be useful for a variety of

applications for cell culture and other applications.

Using the methods described here, PAECs, PAVECs, and PAVICs can be reproducibly seeded

and grown with high viability. A significant amount of trial-and-error was required for this level

of performance, however, and some of the key points are described here. Factors such as cell

seeding density and media exchange frequency were critical in maintaining primary cell viability

and morphology on device. Seeding densities between 2 x 105 – 1 x 10

6 cells/mL coupled with a

media exchange frequency of every 12-16 hours maintained viable primary cells with

appropriate morphologies. Depending on the assay, the cell seeding densities were altered to

vary the duration of culture on device. For example, to demonstrate long-term cell culture,

PAECs were cultured for up to 1 week with an initial seeding density of 2 x 105 cells/mL. For

shorter experiments (e.g., those in which microscopy was performed 24 h after staining), primary

cells were seeded at 5 x 105 - 1 x 10

6 cells/mL, to achieve the desired level of confluence within

24 hours. At densities greater than 2 x 106 cells/mL, cells displayed rounded morphologies with

little spreading, possibly due to overpopulation of the hydrophilic sites and rapid accumulation of

cellular waste products. In all experiments, devices were stored in an incubator in humidified

chambers with no appreciable evaporation.

56

3.3.2 Digital Microfluidic Microscopy, Fixation, Permeabilization, and Staining

As shown in Figure 3-2, DMF devices proved to be a useful platform for microscopic imaging

of primary cells (in this case, using an inverted microscope). For imaging, devices were either

positioned with the bottom plate on the bottom (such that the bottom plate was adjacent to the

objective) as was the case for the images in Figure 3-2, or with top plate on the bottom (such

that the top plate was adjacent to the objective). The capacity to use and flip devices to either

orientation for imaging is a unique property of digital microfluidic devices, which have no open

reservoirs or tubing interconnects that might otherwise interfere. Figure 3-2 shows

representative phase contrast images of PAECs, PAVECs, and PAVICs grown on DMF devices

and for comparison, cells grown on conventional TCPS substrates. As shown, the morphologies

of cultured primary cells were similar on the two surfaces.

Microscopic imaging of cells is often enhanced by staining with fluorescent dyes, which reveals

information about cell state and phenotype. Prior to staining, cells are often fixed to preserve cell

state (by exposure to fixatives such as NBF), and permeabilized to allow for deep penetration by

dyes and other reagents (by exposure to mild surfactants such as Triton X-100). Before and after

these steps and others, the specimen must be repeatedly rinsed as the various reagents can

interfere with each other. Here, as demonstrated in Figure 3-3, we report the first combination of

all of these steps (cell growth, fixation, permeabilization, staining, and rinsing) by DMF. As

shown, at 40x magnification, individual actin stress fibers can be observed, demonstrating the

compatibility of DMF with high-resolution fluorescent microscopy.

57

Figure 3-2: Phase contrast images of PAECs, PAVICs, and PAVECs cultured on a DMF device

(top) and in TCPS flasks (bottom). Scale bar = 200 µm. In the DMF images, the bottom plate is

closest to the objective, and the focus is on the layer of cells on the top plate. The cells are

viewed through the circular optical window between two electrodes on the bottom plate (which

are observable but slightly out of focus).

Figure 3-3: Fluorescent images of PAECs, PAVECs, and PAVICs after fixing, permeabilizing,

and staining on a DMF device. The stains selected for F-actin (FITC-phalloidin, green) and

nuclei (Hoechst, blue). Images were taken at both 10x magnification (top row) and 40x

magnification (bottom row). Scale bar = 200 µm (top row) and 50 µm (bottom row). In these

images, the top plate is closest to the objective.

58

PAECs were cultured on DMF devices and then incubated either with or without TNF-α for 4

hours. Monocytes pre-labeled with Hoechst were then dispensed from reservoirs and delivered to

endothelial cells, which were then rinsed to remove monocytes that did not adhere. As shown in

Figure 3-4, monocytes had greater adhesion to TNF-α-stimulated PAECs compared to non-

stimulated controls, which is consistent with previous studies.9,36-38

These results demonstrate

compatibility of DMF with a fourth cell type (monocytes) and show that primary PAECs

cultured using DMF retain in vivo-like responses to TNF-α. Moreover, this is the first

demonstration of co-culture on a DMF platform. The ability to detect a response with monocytes

(i.e. adhesion) as a result of endothelial cell activation highlights the potential of DMF to

investigate cell-cell interactions.

3.3.3 Digital Microfluidic Monocyte Adhesion Assay

To evaluate the potential for using digital microfluidic systems for co-culture and multistep

assays, we probed its compatibility with endothelial cell/monocyte adhesion experiments.

Monocyte adhesion to endothelial cells is an important initiating event in the inflammatory

process. Endothelial cells are generally activated prior to adhesion, and this state can be induced

by exposure to cytokines such as TNF-α. TNF-α increases monocyte adhesion through

upregulation of EC receptors such as E-selectin,33

intercellular cell adhesion molecule-1 (ICAM-

1)34

and vascular cell adhesion molecule-1 (VCAM-1).34, 35

The assay represented in Figure 3-4

required only a 1.4 µL droplet of reagent and cell suspension for each virtual microwell. In

comparison, macroscale39, 40

and some microchannel-based adhesion assays9 require working

volumes of tens to hundreds of microlitres, such that the DMF system facilitates a 10-100-fold

reduction in reagents used. The capacity to reduce reagent consumption and increase throughput

59

with DMF is desirable in monocyte adhesion assays or other cases in which precious sample or

expensive reagents are used. The potential for combining automated imaging and analysis with

DMF in the future is an attractive vision, as such a system would likely be useful for applications

ranging from basic biology to drug discovery.

3.3.4 Conclusions

We present the first demonstration of primary cell culture using digital microfluidics. A new

mode of “upside-down” culture in virtual microwells was developed to enable primary cell

growth with appropriate morphologies and to decouple the cell growth sites from the digital

microfluidic driving electrodes. Multi-step cell fixation, permeabilization, and staining processes

were demonstrated for the first time on a DMF platform. A monocyte adhesion assay was

performed to demonstrate functionality in DMF-cultured primary ECs and to highlight the co-

culture capabilities of the device. The combination of DMF and primary cell culture/analysis

presented here provides a basis for future studies involving co-culture, high resolution

microscopy, and multiplexed experimentation.

60

Figure 3-4: A monocyte adhesion assay performed on DMF-cultured primary PAECs. (A)

Nuclear-stained (Hoechst, red) THP-1 monocytes adhered to PAECs (calcein AM, green).

Representative images of nuclear-stained monocytes adhered to (B) non-stimulated and (C)

TNF-α-stimulated PAECs. In these images, the top plate is closest to the objective. (D)

Monocytes displayed greater adhesion on TNF-α-stimulated PAECs relative to control non-

stimulated PAECs. Data presented as mean ± standard deviation. *P < 0.05. Scale bar = 200 µm.

61

Table 3-1: Comparison of adherent cell culture using DMF between the new methods reported

here and those previously published.

New Methods

Reported Here

Methods Reported by

Barbulovic-Nad et al.15

Methods Reported by

Lammertyn and

colleagues20,21

Type of cells cultured Primary cells Immortalized cell lines Immortalized cell lines

Pattern of hydrophilic

sites useful for passive

dispensing?

Yes Yes Yes

Location of

hydrophilic sites

Top plate Bottom plate Bottom plate

Hydrophilic site

format

Exposed regions of

ITO surrounded by

Teflon-AF

Spots of adsorbed

fibronectin on a Teflon-

AF surface

Spots of adsorbed poly-

L-lysine on a Teflon AF

surface

Device format for

droplet movement

Right-side up Right-side up Right-side up

Device format for cell

culture

Upside down Right-side up Right-side up

Maximum duration of

cell culture

1 week 2 weeks 3 days

Demonstration of co-

culture

Yes No No

62

Chapter 4. Microgels on-demand

Summary

Three-dimensional (3D) hydrogel particles are enabling technologies in disparate fields ranging

from medical diagnostics to photonics, and they are finding use in fundamental studies in self-

assembly, rheology, and 3D cell culture. Unfortunately, most techniques used for forming 3D

hydrogels are limited to spherical particles and are ‘single pot’ methods in which individual gels

are not addressable after formation. Furthermore, for many applications, it would be useful to be

able to form arrays of gel particles bearing mixtures of constituents and/or are formed from

composites of different gel materials. In response to this challenge, we introduce a digital

microfluidic method for ‘on demand’ formation of arrays of microgels bearing arbitrary

geometries, contents, and shapes. Upon formation of the gels, each particle is individually

addressable for reagent delivery and analysis. We demonstrate the utility of the method for 3D

cell culture and higher order tissue formation by implementing the first sub-microlitre

recapitulation of 3D kidney epithelialization. The new method allows for culture and analysis of

these delicate structures with high success rates relative to conventional techniques relying on

multiwell plates. We anticipate that this platform will provide novel opportunities exploiting

arrays of individually addressable hydrogels of arbitrary shape and size for a wide range of

applications.

63

4.1 Introduction

Precision polymer microgels with complex geometries can support self-assembly for tissue

engineering,83

bar-coding for chemistry,84

and flexible geometric arrangements for forming

photonic crystals.85

A number of strategies relying on microchannels have been developed for

microgel formation including enclosed channels with continuous flows86–88

and two-phase

systems consisting of droplets in a carrier fluid.89,90

Surface tension effects have restricted two-

phase flow systems to the formation of monodisperse spherical, disc, or rod-shaped solids

produced by either photo-initiation or thermal cross-linking at a T-junction or by flow-

focusing.52,53,91

After formation, phase separation requires compatible chemistries to isolate

microgels from the immiscible phase. Recently the Doyle group54,92

improved upon these

techniques, exploiting single-phase stop-flow lithography in the formation of microgels in a

range of geometries. This is important, as arbitrarily shaped microgels (rather than spheres) are

useful for providing novel insights on of the relationship between the microenvironment and cell

fate and behavior.93

The Doyle method has proven particularly useful for UV initiated cross-

linked polymers; however, the method excludes chemically and thermally cross-linked polymer

systems which are important for three-dimensional cell culture. Further, all of the systems

described above are ‘single pot’ methods, such that after formation, the microgels are not

individually addressable.

Motivated by the need to overcome the challenges of polymer compatibility and individual

microgel addressability, we sought to exploit recent developments in reagent dispensing and the

handling of solids on digital microfluidic (DMF) devices for microgel formation. DMF liquid

handling is an emerging alternative to the paradigm of enclosed microchannels.14

This

64

technology facilitates electrostatic manipulation of discrete nano- and micro-litre droplets across

open electrode arrays providing the advantages of single sample addressability, automation, and

parallelization. Variations of this platform have been demonstrated for a broad range of

applications including proteomics,59,94

cell culture and analysis,41,42,72

immunoassays,95,96

chemical synthesis,97,98

and on-chip lasing.64

Recently, the unique geometry of DMF has been

exploited for handling and addressing of three-dimensional solids such as paper discs for genetic

screening,55,56

polymer monoliths for sample extractions,57

and agarose discs for scaffolding

applications.59,99

Here we recognized the potential for DMF to address obstacles to microgel

formation: (1) on-demand hydrogel formation, (2) flexible hydrogel geometries, (3) single

hydrogel addressability, and (4) compatibility with UV, chemical, or thermally cross-linked

polymer systems. Finally, we propose that the combination of digital microfluidics and hydrogels

may represent a useful new tool for three dimensional cell culture, an important emerging

technique for in vitro biology44

that is not widely used because of experimental complexity

(diffusion into 3D matrices is slow and inconvenient for automation) and hydrogel fragility (3D

matrix materials break down upon repeated handling and fluid exchange).

We introduce here the first method for forming “microgels on-demand,” in which individually

addressable three-dimensional (3D) gel structures of varying sizes, shapes, and compositions can

be formed in situ. These advances have the potential to overcome current challenges in complex

microgel formation and addressability, with particular emphasis on 3D cell culture for drug-

screening with improved physiological relevance.

65

4.2 Methods

4.2.1 Reagents


Canada) or Fisher Scientific Canada (Ottawa, ON, Canada), antibodies, fluorescent dyes, and,

cell media components were from Invitrogen/Life Technologies (Burlington, ON, Canada), and



4.2.2 DMF device fabrication

Digital microfluidic devices were fabricated using standard photolithography and metal etching

as detailed previously.42

The bottom-plate device design featured an array of 2.2 mm × 2.2 mm

chromium actuation electrodes and also included an array of five 1 mm diameter optical

windows (i.e., circular regions free from chromium) with 9 mm between each window. Each

window straddled the interface between two actuation electrodes. DMF device top-plates bearing

hydrophilic sites were formed by performing a Teflon liftoff procedure on ITO coated glass

substrates as detailed in chapter 242

The sites were 1-2 mm in diameter and were either circular,

star, heart, or diamond in geometry. The diameter for non-circular shapes was defined as the

diameter of the smallest circle that would enclose the feature.

4.2.3 DMF device assembly and operation

Digital microfluidic devices were assembled with an ITO–glass top plate and a chromium-glass

bottom plate as shown in

Figure 4-1. The two plates were joined by stacking two, three, or four layers of double-sided

tape (each layer ~80 µm), and were aligned such that the edge of the top plate was adjacent to the

66

outer-edges of the reservoir electrodes on the bottom plate. Care was taken to align top and

bottom plate features vertically (windows on the bottom plate and hydrophilic sites on the top

plate). A driving potential of 120 VRMS was generated by amplifying the square wave output of a

function generator (Agilent Technologies, Santa Clara, CA) operating at 10 kHz. Reagents were

loaded and dispensed, moved, and merged as described previously.42

In all DMF experiments,

reagent solutions were supplemented with 0.02% Pluronics F68 except for sol-state Geltrex that

was supplemented with 0.02% Pluronics F127.74

Agarose was not supplemented with Pluronics.

4.2.4 Hydrogel pillar formation and addressing

Devices bearing droplets were imaged with a CCD camera (Basler, Ahrensburg, Germany)

mounted on a fluorescence equipped stereomicroscope (Leica, Wetzlar, Germany). Geltrex was

prepared by 1:1 dilution in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with

10% FBS and 0.02% F127. The solution was maintained on ice until device loading. Low gelling

temperature agarose hydrogels were prepared as 1-6% w/v solutions in DI water by microwaving

the solution for 30 seconds prior to device loading. In some cases sol-state hydrogel solutions

were supplemented with 10 µM fluorescein or with a suspension of fluorescently labelled (green,

yellow, or red) 10 µm dia. microspheres. To form hydrogel pillars, 5 µL of sol-state gel solutions

were loaded into device reservoirs, then one or two droplets were actively dispensed and then

manipulated across the patterned hydrophilic sites where sub-droplets were generated by

hydrophobic-hydrophilic interactions.42

Devices containing Geltrex and agarose sub-droplets

were cross-linked to form pillars by incubation for 1-4 hours at 37 ºC in a humidified chamber,

or at room temperature, respectively. After formation, individual reagents (e.g., fixatives,

permeabilizers, dyes, etc., as described below) were delivered to individual gel pillars by loading

67

Figure 4-1: Digital microfluidic device geometry. (A) Exploded view of device, comprising a

bottom plate with patterned electrodes and a top plate bearing patterned hydrophilic sites. (B)

Side-view of device, not to scale. (C) Schematic depicting principle of device operation.

68

the appropriate mixture into a reservoir, actively dispensing a 0.4 L to 2.2 L droplet onto the

array of electrodes, and then passing the droplet across the pillars (passively exchanging the

contents of the pillar).

4.2.5 Diffusion analysis and modelling

Hydrogel pillars were formed from Geltrex and agarose in 1.5 mm dia. circular virtual

microwells. After two hours of cross-linking at room temperature in a humidified flask, a droplet

of either fluorescein (10 µM in DI water) or fluorescein isothiocyanate conjugated dextran (4

kDa or 40 kDa, 25 mg/mL in DI water) was passed across each hydrogel in a temperature-

controlled imaging suite at 23 ºC. Each condition was repeated in triplicate. Fluorescent images

were recorded at five frames per second (FPS) for up to three hours. Fifty frames were selected

for analysis from each experiment between the onset of diffusion and saturation. Image analysis

was performed in ImageJ (http://rsb.info.nih.gov/ij/) by integrating the pixel intensity in a ~1.3

mm dia. circular region of interest (ROI) within the hydrogelfor each frame over the

experimental observation time. The apparent diffusion coefficient (Da) was determined by an

adaptation of the method reported by Axelrod et al.100

for fluorescence photobleaching recovery.

This model assumes no source density, uniform pore size, and no evaporation. Briefly, integrated

pixel intensities were normalized to the initial fluorscence intensity of the carrier droplet. These

were then plotted with respect to time and the plateau intensity (A) was determined. A

flourescence saturation half-point (τ1/2) was defined as the time required for the normalized

intensity to reach ½A. The time constant for diffusion (τ) was determined by:

69

2/1

)2

1ln(

Eqn. 4.1

An empirical curve was fit to the experimental data:

))exp(1()(

tAxtf Eqn. 4.2

And the apparent diffusion coefficient was set to, where r is the radius of the hydrogel:

2/1

2

4

88.0

rDa Eqn. 4.3

A numerical simulation was performed using COMSOL Multiphysics (http://www.comsol.com/).

A two-dimensional diffusion model was implemented with boundary conditions set to a constant

concentration and the respective diffusion coefficients from Eq. 3. These experiments were

performed with ~160 µm spacers between plates.

4.2.6 Composite hydrogel formation

To build composite hydrogels a cross-linked microgel was formed initially as described above

from either Geltrex or agarose. Secondary hydrogel structures were then formed by manipulating

sol-phase hydrogel droplets to surround the initial structures. These were then cross-linked by

room temperature incubation and imaged under UV illumination with a digital camera. These

experiments were performed ~160 µm spacers between plates.

70

4.2.7 Cell Culture

Marbin Darby canine kidney (MDCK) epithelial cells were kindly provided by Dr. N. Tufenkji

(McGill University). MDCKs were cultured in DMEM supplemented with 10% fetal bovine

serum (FBS), 100 U∙mL-1

penicillin, and 100 µg∙mL-1

streptomycin.

4.2.8 Cell viability and cell distribution

MDCK cells were prepared at 1 106 cells∙mL

-1 in culture media then diluted by 50% in 4ºC

Geltrex solution. 5 µL aliquots of this suspension were loaded and droplets were actively

dispensed from reservoirs and then driven across 1.5 mm circular hydrophilic sites to form

hydrogel pillars bearing suspended cells. Each device was then inverted, placed within a petri

dish containing kimwipes saturated with DI, and incubated at 37 °C in a humidified incubator

containing 5% CO2 for two hours. Cells were then stained by manipulating PBS droplets

supplemented with Live/Dead reagents (Life Technologies) and 5 µg/mL Hoeschst 33342 to

exchange the solution with the pillars. Cells in the hydrogel pillar were imaged by confocal

microscopy (Zeiss LSM700, Carl Zeiss, Toronto, Canada) at 6 µm increments along the z-axis.

Viability after cross-linking was enumerating the number of live (green) fluorescent cells, and

dead (red) fluorescent cells. Viable spheroids were evaluated on days two through four by light

microscopy, where viable spheroids have obvious spheroid shapes and non-viable spheroids

appear disorganized and/or blebbed. Cell distribution was determined by recording the z position

of the largest diameter portion of each individual nucleus throughout the hydrogel. Image

analysis was performed using Zen Light Edition software (Carl Zeiss, Toronto, Canada). These

experiments were performed with ~240 µm spacers between plates.

71

4.2.9 Epithelialization experiments

MDCK cells were seeded in pillars on devices as described above. Cells seeded on separate

devices were fixed at 24, 48, and 96 hours after initial seeding by passing three droplets of 10%

v/v Histochoice Tissue Fixative in DI water to each hydrogel pillar then incubating for 30

minutes at room temperature. Cells were then permeabilized by passing three droplets of

permeabilization solution (PS, 0.5% Tween 20 diluted in PBS) across each hydrogel pillar then

incubating for 90 minutes at room temperature. Prior to staining, three droplets of PS were

driven across the hydrogel with 10 minute incubation at room temperature between each droplet.

Each gel pillar was addressed with a droplet containing staining reagents and then incubated at

room temperature for up to 3 hours in the dark. Prior to imaging, three droplets of PS were

passed across each gel pillar with 10 minute incubation at room temperature between each

droplet. Spheroid diameter was determined by light microscopy and cell number per spheroid

was determined by enumerating nuclei per spheroid by epifluorescent microscopy. Confocal

microscopy was used to image cells stained with Alexa Fluor 488® phalloidin and Hoescht

33342. These experiments were performed with ~240 µm spacers between plates.

4.2.10 Comparison to conventional fluid handling

MDCK spheroids were formed in 3D culture as described above on DMF or in 96-well plates.

On day four of culture ten randomly selected spheroids for each condition were imaged by light

microscopy, followed by reagent exchange. For DMF this comprised manipulation of five

separate 1.2 µL droplets of PBS across the microgel containing spheroids. For 96-well plates, a

Biomek FX system (Beckman Coulter, Inc., Fullerton, CA, USA) was used to deliver 100 µL of

72

PBS and then remove 100 µL of PBS at a rate of 50 µL/s from each well five times. Spheroids

were then re-imaged post manipulation and compared to spheroids pre-manipulation by a

measure of circularity using ImageJ to indicate the level of spheroid deformation. Spheroids

were identified by automated microscopy homing (to pre-selected x, y coordinates) and the use

of other local structures as fiduciary marks. Circularity was determined by drawing an ROI along

the mid-point of each spheroid (before and after manipulation) and calculated as:

24

perimeter

areayciruclarit Eqn. 4.4

Deformation was determined as the absolute percent change in circularity before and after

manipulation, where 100% would indicate complete spheroid disintegration. These experiments

were performed with ~240 µm spacers between plates.

4.3 Results and discussion

4.3.1 Microgels on-demand

To form microgels on-demand, droplets of sol-phase hydrogel material are manipulated across

hydrophilic sites on a DMF device (

Figure 4-1,Figure 4-2), where sub-droplets are generated by line-pinning. This technique has

been used previously for non-gelling fluids (known in DMF as “passive dispensing”42

and in

non-DMF methods as “surface energy traps”101

), but this is the first application to exploit this

technique for hydrogels. As shown in Figure 4-2, in this method, sub-droplets of sol-form

hydrogel materials deposited onto hydrophilic sites can be thermally cross-linked to form

73

Figure 4-2: Microgels on-demand. (A-H) Frames from a movie (top) and side-view schematic

(bottom) depicting a sol-state hydrogel droplet containing a fluorescent dye actively dispensed

from a reservoir (A) and then electrostatically manipulated to a patterned hydrophilic site (B &

C). A second droplet is then actively dispensed (D) and passed across the hydrophilic site (E-H).

Scale bars = 2 mm. When the sol-state droplet is passed across the hydrophilic site a sub-droplet

is generated in the shape of the hydrophilic site. Upon crosslinking, each droplet forms a solid

gel pillar. Various geometries can be generated and visualized with epifluorescent

stereomicroscopy (I & J) or confocal microscopy (K & L). A movie depicting the formation of

an array of different microgel shapes can be found online in the supplementary information.

Scale bars = 1 mm.

74

microgel pillars. The two materials used here were Geltrex (a reconstituted basement membrane

complex of extracellular matrix (ECM) proteins102

that remains in sol-phase at temperatures

below 4°C) and agarose (a gel that remains in sol-phase at temperatures above 42°C), but we

propose that similar methods should be applicable to a wide range of hydrogel materials. In the

only previous work that we are aware of describing the use of hydrogels on DMF, gel discs were

formed off-device then manually positioned on device (without passive dispensing).59,99

Though

useful, the previously described method is limited to circular structures, cannot be implemented

on-demand, and is incompatible with soft gel materials such as those that are based on collagen

(like Geltrex). We found that the microgel on-demand method could repeatedly produce

columnar hydrogels formed from both agarose and Geltrex with dimensions ranging from 1000-

2000 µm diameter and 75-225 µm in height (~60-700 nL) with volumetric precision varying

from 0.3% to 8.1% (Figure 4-3).

The formation of gels with arbitrary shapes has previously only been possible using UV cross-

linkable hydrogels; however, UV initiators are often cytotoxic and therefore are typically not

ideal for cell applications.54

Here, we report that by simply altering the shape of the hydrophilic

site for passive dispensing, star-, heart- triangle-, and diamond-shaped microgels were readily

formed using the microgel on-demand technique (Figure 4-2,I-J, Figure 4-4). We further

confirmed that these microgels were conformal through the vertical axis by confocal microscopy

(Figure 4-2,K-L). Given the extensive evidence in the literature demonstrating the effects of two-

dimensional geometry on cell function and the emerging evidence that 3D microenvironmental

geometry plays a role in tissue morphogenesis,93

we propose that the microgel on demand

method will be a useful new tool for probing the role of microenvironment geometry on cell and

75

Figure 4-3: Precision of microgel on-demand formation and reagent exchange. Frames from a

movie (A-D) depicting sol-state Geltrex being passively dispensed to form a sub-droplet. Scale

bar = 2 mm. The diameters of sol-state Geltrex were measured by brightfield microscopy and

compared to the diameters of circular hydrophilic sites (E). The asterisk indicates the single

condition tested for which sub-droplets failed to form. Experiments were performed in triplicate

and error bars indicate ±1 S.D.

76

Figure 4-4: Formation of microgels in varied geometries. Frames from movies depicting the

formation of star (A-D), triangle (E-H), and diamond-shaped microgels (I-L). Microgel density

can be increased by adjusting the pitch of patterned hydrophilic sites on the device top-plate (M).

A movie depicting the formation of an array of microgels in different shapes can be found in the

online supplementary information. Scale bars = 2 mm.

77

tissue behavior and function.93,103

Finally, microgels (with sufficient rigidity) formed on demand

are accessible for off-chip handling and analysis. Agarose gels prepared in this manner were

imaged by environmental scanning electron microscopy (ESEM) (Figure 4-5), revealing that

they possess the honeycomb structure typical of agarose microgels formed by other means.104

In

contrast, Geltrex microgels were not sufficiently rigid to maintain their form after top-plate

removal.

4.3.2 Reagent exchange and diffusion into hydrogels

A key benefit of DMF liquid handling is the independent addressability of electrodes and thereby

individual droplets. This feature allows for the targeted delivery of reagents, an important

attribute for the maintenance of microgel hydration and cell viability, in the case of three-

dimensional cell culture, through media exchange. As shown in Figure 4-6A-D, droplets of

reagents were individually delivered to (and removed from) microgels by applying an

appropriate sequence of driving potentials. This dispensing mechanism is reproducible and

precise, ensuring that each passage delivers equal amounts of fresh reagent with volumetric

precision varying from 0.29% to 3.29% (Figure 4-7).

To examine diffusion into DMF-generated microgels we used fluorescein (0.3 kDa), 4-, and 40-

kDa FITC-dextrans as surrogates for the small molecules, growth factors, and cytokines often

used in cell culture and stimulation.105

Specifically, droplets containing these tracers were

78

Figure 4-5: Off-chip microgel analysis. Agarose (5% w/v) hydrogels were formed on device

then allowed to crosslink at ambient temperature. Top-plates were removed to expose the

hydrogel allowing for imaging by environmental scanning electron microscopy.

79

delivered to Geltrex microgels formed on-demand and diffusion was tracked by fluorescence

imaging. Upon delivery of a droplet, a film of tracer was observed to form around the microgel

and diffuse towards the centre. As shown in Figure 4-6E-F, the three tracers had apparent

diffusion coefficients (Da) in Geltrex of Da = 10×10-10

m2/s (fluorescein), 7.2×10

-11 m

2/s (4-kDa

FITC-dextran), and 3.1×10-11

m2/s (40-kDa FITC-dextran), respectively. For comparison,

diffusion was modeled numerically, which was found to be consistent with the experimental

results (Figure 4-6E). The deviation between modeled and experimental results could be

attributed to the depleting level of Co within the absorbed ring of tracer over time. Further, the

experimental values are consistent with the limited number of diffusion coefficients, D, reported

in the literature for similar (but not identical) systems: fluorescein in water106

(D = 4.9×10-10

m2/s) and matrigel107 (D = 4.2×10-10

m2/s), 8-kDa FITC-dextran in agarose108

(D = 8×10-11

m2/s),

20 kDa FITC-dextran in water109 (D = 8×10-11

m2/s) and agarose105 (D = 4.2×10-11

m2/s). The Da

values determined here suggest that delivery of small molecules, growth factors, or cytokines for

hydrogel based cell culture will be fast, requiring only ~minutes to reach cells embedded within

the gels (in comparison to ~hours for macro-scale 3D cell culture in multiwell plates).

4.3.3 Pitch, Multi-Component Arrays, and Composite Microgels

In most of the data described herein, the spacing between gels (or “pitch”) was 9 mm, which

enabled the formation of five gels at a time on the ~microscope slide sized substrates used here.

But the spatial density of hydrogel structures can be readily increased by reduced pitch and size

of individual hydrogel structures (Figure 4-8A). In addition, the new technique facilitates the

formation of combinatorial arrays of microgels containing unique constituents. To highlight this

80

Figure 4-6: Reagent exchange in hydrogel pillars. (A-D) Frames from a movie depicting a

droplet of fluorescein being electrostatically manipulated across a (transparent) microgel pillar

and subsequent diffusion of fluorescein into the microgel. Scale bars = 2 mm. (E) Experimental

(red dots), empirical fit (dashed blue line), and simulation (solid black line) of diffusion profiles

for fluorescein into Geltrex microgels. The inset shows a fluoresecent image (left) and heat-map

simulation of concentration (right) of fluorescein at the half-saturation point (1/2). (F) Apparent

diffusion (Da) coefficients measured for fluorescein, 4 kDa FITC-dextran, and 40 kDa FITC-

Dextran into Geltrex. Error bars indicate 1 S.D.

81

Figure 4-7: Precision of reagent exchange across microgels. Frames from a movie (A-D)

depicting a droplet of reagent passed across a microgel. The diameters of dispensed droplets

were measured by brightfield microscopy and were compared to the diameters of circular

hydrophilic sites (E). The asterisk indicates the single condition tested for which sub-droplet

failed to form. Experiments were performed in triplicate and error bars indicate ±1 S.D.

82

capability, we generated arrays of hydrogel mixtures of red, yellow, and green constituents,

forming a gradient of compositions (Figure 4-8B). These arrays were generated combinatorially

on device from three stock sol-phase solutions (each containing only one colour of microsphere)

through a series of droplet merging, mixing, and splitting steps. We propose that this technique

may be useful in the future to screen arrays of different hydrogels compositions and multiple cell

types for studies in tissue engineering and cell-ECM interactions.110,111

Finally, the technique can

be used to individually address microgels to form composite materials by forming a first

structure from either Geltrex or agarose, followed by the formation of a secondary hydrogel

structure engulfing the initial structure (Figure 4-8C-E). Composite microgel structures have

been reported previously, but they have been limited to spherical geometries.112

We propose that

the new capacity to form arbritrarily shaped composite gels may be useful for forming unique

environments for multi-scaffold chemistries and cell culture.110,113

4.3.4 DMF recapitulation of epithelialization

With the ability to form microgels on-demand by DMF and to address them independently for

reagent exchange, we tested this system for three dimensional cell culture using the well-

characterized model of kidney epithelialization.44,45,114

MDCK cells were suspended in sol-phase

Geltrex and this suspension was used to form microgels on-demand, as above (Figure 4-9A).

Cells cultured in microgels remained viable (~100%, data not shown) for up to 5 days, with

delivery of fresh media droplets at 24-h intervals. The effects of electrostatic actuation on cell

health were not explicitly evaluated here, but previous studies with 2D adherent or suspension

cell culture have reported no or negligible effects on cell viability/morphology67,72

or gene

expression115

when compared with non-actuated cells (because the electrical field drops across

83

Figure 4-8: Pitch, multicomponent arrays, and composite microgels. 32-plex arrays of microgels

were formed from Geltrex supplemented with fluorescein for visualization (A). Combinatorial

arrays of Geltrex microgels were formed on device containing mixtures of red, yellow, and/or

green microspheres (B). Gradient bars indicate the percent compositions of each respective

microsphere, from 100% red at the left and 100% green at the right. Composite microgels

containing fluorescent microspheres were formed with inner agarose and outer Geltrex layers (C:

green filter, D: red filter, E: composite image). Scale bars = 2 mm.

84

the insulating layer rather than droplets containing cells). As shown in Figure 4-9B and Figure

4-10, the cells evaluated here were found to be suspended within the hydrogel matrix with a

slight bias to the bottom of the device, with no cells adhered to either top or bottom plates.

MDCKs are known to form hollow spheroid structures when cultured in collagen or matrigel

matrices, making it possible to study epithelialization and primitive tissue formation in vitro.45

To evaluate whether cells cultured in microgel pillars on DMF could recapitulate this model,

MDCKs were maintained on-device with daily media exchange for one through four days. On

each day microgels were fixed, permeabilized and stained for actin and nuclei (all steps

implemented as in Figure 4-6, allowing for delivery of reagents in minutes). The inherent

addressability of DMF makes this process straightforward, allowing for one gel to be stained and

imaged on day one (while maintaining cell growth in the other gels), a second gel to be stained

and imaged on day two, and so on. Cells formed multicellular clusters in DMF-microgels over

the first 72 hours (Figure 4-9C-F) with a visible lumen observed after 96 hours. Nuclei were

distributed around well-formed lumina with actin accumulation at the luminal edge, an indication

of cellular polarization. The sizes of spheroids cultured within sub-microlitre gel pillars were

similar to those grown in 96-well plates in 100 L aliquots (Figure 4-11).

The results in Figure 4-9 represent the first demonstration of MDCK spheroid formation in sub-

microlitre hydrogels (using microfluidics or any other format). Note that these spheroids are

higher-order structures that form spontaneously as a result of cell directed assembly; these

structures are quite distinct from those in which cells are grown on pre-formed hydrogel targets

85

Figure 4-9: Higher order tissue formation and handling in microgel pillars. Spheroids are

generated five-plex in microgels on DMF (A). The inset highlights one microgel containing

spheroids. Confocal microscopy image stack of MDCK cells in a Geltrex pillar demonstrates cell

distribution throughout the z-axis (B). Images depicting MDCK spheroid formation in a gel pillar

over four days in culture (C-F). Cells were stained for actin with phalloidin (green) and nuclei

(blue). On day four lumen formation was observed. Hydrogels bearing 4-day MDCK spheroids

were subjected to media exchange five consecutive times either by automated pipetting or DMF.

Representative bright-field images of spheroids pre- and post-manipulation for both systems (G-

J). Scale bars = 20 µm. Graph of spheroid deformation for five trials evaluating 10 spheroids

each using robotic pipetting into a multiwell plate or DMF (K). The deformation value of each

spheroid is indicated by a blue (DMF) or red (conventional reagent delivery) dot with black bars

representing the mean deformation of each trial. Bar graphs show overall deformation averages

across the five trials for each respective system. Error bars show 1 S.D. In three of the trials,

identified by asterisks, the conventional method dislodged the hydrogels from the well, resulting

in material failure.

86

Figure 4-10: Z-axis distribution of cells in microgels. Graph of distribution of cells in the z-axis

for the data in Figure 4-9B. The number of cells was enumerated at each image plane (~6 µm

thick).

Figure 4-11: Spheroid size. Comparison of spheroid size for three-dimensional MDCK culture in

microscale (DMF, white bars) and macroscale (96-well plate, gray bars).

87

or molds.116,117

One reason for the relative dearth of this type of work in the literature is that

higher order tissue structures are very fragile. The simple act of delivering reagents to 3D cell

structures is known to cause morphological damage51,118

; this is particularly problematic for cells

grown in hydrogel scaffolds with low viscoelastic storage modulus (~10 Pa for 50% Matrigel119

).

We hypothesized that digital microfluidics, which has been previously reported to be useful for

gentle handling of weakly adhered apoptotic cells,72

might also be useful for non-disruptive

reagent delivery to cell constructs grown in 3D microgels.

To characterize and compare the damage induced to MDCK spheroids by reagent exchange, we

performed assays in which day-4 spheroids (cultured either in microgels on DMF devices or in

wells on microtitre plates) were imaged and then five aliquots of PBS were added and removed.

The spheroids were then imaged again, and deformation was determined by calculating the

change in circularity of each spheroid’s two-dimensional projection imaged by light microscopy

(Figure 4-9G-J). As shown in Figure 4-9K, in five separate trials (evaluating 10 spheroids in

each trial), DMF handling of hydrogel bearing spheroids resulted in a nearly ten-fold reduction

in deformation when compared to conventional fluid delivery to cells in mutliwell plates (DMF

7.6%.and conventional 75.9%). Of particular note are three trials in which conventional handling

of the microgels resulted in complete hydrogel displacement (quantified as 100% deformation),

in which spheroids were disrupted to the extent that they could no longer be identified. This

evidence supports the hypothesis that DMF is superior to conventional techniques for gentle

handling of microgels, an important factor in terms of the cost-benefit of 3D cell culture.

88

4.3.5 Conclusion

Digital microfluidic (DMF) microgels on-demand is a new tool for the formation of microgels in

a range of geometries and compositions. Importantly, this is the first method capable of forming

complex and composite hydrogel geometries from thermally cross-linked hydrogels, with

individual addressability of microgel structures. With demonstrated utility for three-dimensional

cell culture, we anticipate this technology will enable future microgel studies across a range of

applications in chemistry, biology, physics, and beyond.

89

Chapter 5. Cardiac microgels

Summary

Three-dimensional (3D) cell culture is attractive because of the ability to better reproduce in vivo

biology in vitro. Unfortunately, 3D cell culture techniques are much less common than their 2D

analogues because the fragility of matrix materials makes them difficult to manipulate using

conventional tools, and because the reagents are expensive. Moreover, most techniques used to

form 3D cell constructs form them and address them in bulk dispersions; ideally, methods could

be developed to evaluate and address each 3D tissue one-at-a-time. Here we describe the

application of ‘microgels on-demand’, a flexible method for micro-scale hydrogel formation for

3D cell culture and analysis, for the culture of functional cardiomyocyte (CM) microtissues. In

this technique, the unique fluid handling capabilities of digital microfluidics (DMF) is leveraged

to generate, address, and maintain sub-microlitre gel pillars that support the growth of 3D

neonatal rat CM structures. Over the course of 5 days, these ‘cardiac microgels’ initiate

spontaneous beating and are independently addressable and responsive when stimulated with

epinephrine. Further we report a new label-free method for analyzing CM activity which may

someday facilitate integration with high throughput processing for screening experiments. This

DMF based technique supports cardiac microgel formation and we anticipate access to this

technology to enable improved biological readouts from in vitro assays at significantly reduced

costs.

90

5.1 Introduction

Cardiac tissue presents a complex three-dimensional environment composed primarily of

contractile muscle cells and fibroblasts. The interaction of cardiomyocytes (CMs) with the

surrounding extracellular matrix is critical for the formation of anchor points that transduce the

mechanical signals needed to promote CM formation of myofibrils and sarcomeres. Cultured

primary CMs are important for the study of cardiac function in vitro. Since the initial isolation of

neonatal rat CMs by Harary and Farley,120

these cells have been well characterized in terms of

their morphological, biochemical, and electrophysiological properties and are commonly used to

evaluate cardiac toxicities of drugs. To date, the majority of studies have focused on CMs

cultured on planar two-dimensional (2D) plastic or glass surfaces. Though the data generated

from these studies has proven invaluable, there remains a significant disparity between 2D in

vitro model systems and observed in vivo responses. For this reason microtissue models for in

vitro assays are being developed to bridge the gap between traditional two-dimensional cell

culture systems and in vivo biology.121,122

Hydrogel-based 3D cell culture is rapidly becoming an important tool in life-science research.44

Hydrogel materials can be derived from inert or animal sources providing for customizable

microenvironments to elucidate or direct cell function and behaviour. These systems have

allowed for significant strides in cell biology through the reestablishment of critical

microenvironmental factors, particularly cell-cell and cell-matrix interactions. To improve the

reliability of data from drug screens and avoid the pursuit of in vivo follow-up studies on false-

positive targets, 3D cell culture presents a potentially ideal system to balance cost and biological

relevance. Unfortunately, hydrogel culture methods remain under-utilized in part because of high

91

reagent costs and challenges in the manipulation and handling of delicate hydrogel materials.

A number of strategies relying on microchannels have been developed to address the challenges

of working with 3D cell structures in hydrogels,47

including enclosed channels with continuous

flows86,87

and two-phase systems consisting of droplets in a carrier fluid.89,90

Microfluidics is

useful for this goal, as it provides the ability to manipulate sub-microlitre volumes of liquid

thereby reducing reagent consumption. Devices used for 3D cell culture and handling have

ranged in complexity from systems with integrated valves48

and off-chip pumps52,123

to passive

perfusion systems50,51

exploiting gravity and surface tension forces to drive reagent exchange.

Microchannel devices have been demonstrated for 3D culture of primary CMs, to pattern

CMs,124

to form two-dimensional culture systems on hydrogel coated surfaces,125–127

or to enable

3D spheroid culture.128

Though these examples successfully form functional cardiac microtissues

in their respective formats, they are ‘single pot’ systems, and inherently lack the ability to

individually address each microtissue during experimentation.

To address this challenge we recently developed a digital microfluidic method (DMF) called

‘microgels on-demand’, introduced in Chapter 4, that allows for the formation of microgels in

customizable geometries from a broad range of hydrogel materials. This method was

successfully implemented in the recapitulation of a model of kidney epithelialization. DMF

based biological assays facilitate the manipulation of sub-microlitre volumes of reagent across

arrays of electrodes and have been used for a broad range of cell based assays with immortalized

and primary cells cultured in both 2D and 3D formats (several examples of which are presented

in chapters 2, 3, and 4).41,58,67,72

While DMF actuation has been demonstrated to exert little if

92

any transcriptional effect on cells,115

the small volumes required (< 1 µL) are particularly

attractive for primary cell culture because of the inherently limited amounts of available material.

Contraction frequency is well-known property that is used classically to evaluate CM function –

with an emphasis in microtissue models to achieve physiological rates of contraction in

vitro.121,122,124,126

Unfortunately, the measurement of contraction frequency is significantly more

challenging in 3D culture than in 2D culture. Moreover, traditional methods extrapolate the

behaviour of tissue constructs from the morophological changes in single cells. To circumvent

these challenges, we developed a whole frame method to quickly ascertain ‘CM activity’ within

3D constructs. This metric can be correlated with CM beat frequency, providing a rapid, simple

and novel high-level analysis technique in the determination of 3D CM activity. We anticipate

the methods reported here, if widely applied, will enable laboratories to improve efficiency in

screening CM based culture systems, while reducing the amount of material required (and thus

the numbers of animals to be sacrificed).

5.2 Methods

5.2.1 Reagents


Canada) or Fisher Scientific Canada (Ottawa, ON, Canada), antibodies, fluorescent dyes, and,

cell media components were from Invitrogen/Life Technologies (Burlington, ON, Canada), and



93

5.2.2 DMF device fabrication

Digital microfluidic devices were fabricated using standard photolithography and metal etching

as detailed previously.42

The bottom-plate device design featured an array of 2.2 mm × 2.2 mm

chromium actuation electrodes and also included an array of four 1.5 mm diameter optical

windows (i.e., circular regions free from chromium) with 9 mm between each window. Each

window straddled the interface between two actuation electrodes. DMF device top-plates bearing

1.5 mm dia. circular hydrophilic sites were formed by performing a Teflon liftoff procedure on

ITO coated glass substrates as detailed in Chapter 2.42

5.2.3 DMF device assembly and operation

Prior to experiments, digital microfluidic top and bottom plates were sterilized in 70% ethanol

for 10 minutes. Excess ethanol was shaken off and devices were permitted to air dry for 30

minutes within a biosafety cabinet. DMF devices were assembled with an ITO–glass top plate

and a chromium-glass bottom plate. The two plates were joined by three layers of double-sided

tape (each layer ~80 µm), and aligned such that the edge of the top plate was adjacent to the

outer-edges of the reservoir electrodes on the bottom plate. Care was taken to align top and

bottom plate features vertically (windows on the bottom plate and hydrophilic sites on the top

plate). An open-source, automated DMF actuation system called “DropBot” (described in detail

elsewhere129

) was used to program and manage the application of driving potentials of 120-140

VRMS generated by amplifying the sine wave output of a built-in function generator operating at

10 kHz. Reagents were loaded and dispensed, moved, and merged as described previously.42

In

all DMF experiments, reagent solutions were supplemented with 0.02% Pluronics F68 except for

sol-state Geltrex that was supplemented with 0.02% Pluronics F127.74

Imaging during droplet

manipulation was performed with a built-in webcam.129

94

5.2.4 Cell culture and cardiac microgel formation/addressing

CMs were isolated from neonatal (24-48 hour old) Sprague-Dawley rat hearts as described

previously130

and cultured in DMEM supplemented with 10% FBS, 1% HEPES, 100 U/mL

penicillin-streptomycin, 0.02 U/mL insulin, and 5 µg/mL vitamin C. For microscale 3D cell

culture experiments, cell suspensions (5-20×106 cells/mL) in sol-state gel were prepared in a

50% Geltrex solution in CM media. To form microgels, an 8 µL aliquot of sol-state gel/cell

suspension was loaded into device reservoirs. The volume of the entire media reservoir was

dispensed onto the platform and then translated across the patterned hydrophilic sites to generate

sub-droplets by hydrophobic-hydrophilic interactions.42

Devices were inverted then placed into a

Petri dish in a 150 cm2

tissue culture flask with re-closable lid containing DI water. The flask

was then incubated for 4 hours at 37 ºC in a humidified chamber to allow for cross-linking. After

cross-linking, devices were returned to upright state, reconnected to DropBot, and fresh media

was delivered to the microgels by single droplet passive exchange. Subsequent media exchanges

were performed every 24 hours for up to five days (with devices stored inverted in the incubator

between exchanges). For macroscale 2D cell culture experiments, cells were seeded at 50,000

cells per well on a 96-well plate that had been coated with Geltrex (1:30 dilution in CM media).

Media was exchange at 24 hour intervals after seeding for up to five days.

5.2.5 Cardiomyocyte treatment

Media was exchanged by passive dispensing and cells were maintained in an incubator for 30

minutes prior to collecting a movie of CM activity for 20 seconds (30 frames per second, fps) at

4×, 10×, or 20× magnification with a brightfield microscope (Motic AE31, Japan) equipped with

a black and white camera (Basler A640, Exton, PA) controlled by a custom Labview program

95

(National Instruments, Toronto, Canada). Cells were then treated with either the stimulant

epinephrine (10 µg/mL in CM media) or the fixative histochoice (10% in DI water) by passive

reagent exchange. In the case of epinephrine stimulation, cells were incubated for 20 minutes

followed by imaging as described above. Cells were then washed with fresh media, incubated for

20 minutes, then fixed. In certain experiments, cells were fixed in histochoice (10% in DI), then

imaged for 30 minutes (30 fps) at 10× magnification. Cells cultured in 2D macroscale culture

were treated with similar conditions.

5.2.6 Single cell analysis of cardiomyocyte activity

Cells were imaged by light microscopy at 10× and 20× magnification. Single cell function was

determined by evaluating cell body displacement, contraction frequency, and contraction length

from videos of beating tissues. Specifically, ImageJ was used to determine the regions where

cells were contracted. Subsequent relaxation was determined by cell movement relative to the

original region of interest. The distance of the contraction was measured as the greatest distance

travelled by any single edge of the cell. Twenty cells were evaluated in three independent micro

gels to determine overall functionality.

5.2.7 Cardiac activity coefficient (CAC) for fast analysis

Cells were imaged by light microscopy at 10× and 20× magnification. A custom Matlab

(Mathworks, Nattick, MA) script was written to evaluate changes in pixel intensity across image

sequences. With a maximum observed beat rate of less than 120 BPM, we selected a sampling

frequency to avoid aliasing using the Nyquist criterion:

cs ff 2 Eqn. 5.1

96

where fs is the sampling frequency and fc is the cutoff frequency. To meet this criterion, we

analyzed image frames at a frequency of 6 fps. In the analysis, the Matlab script determined the

mean change in 8 bit pixel intensity between every sample frame (every 0.167 s); we termed this

the cardiac activity coefficient (CAC). This is described by the following equation:

N

II

CAC

N

i

ii

1

Eqn. 5.2

Where I is the image intensity and N is the total number of images. In certain experiments this

was presented as a function of time. In others, the mean pixel intensity over a twenty second

video at 0.167 intervals was determined. These were either collected as a sum of pixel change

over the course of a twenty second sequence, or the average over the course of a longer

experiment. High and low thresholds were determined by finding maximum and minimum

values in each condition. These thresholds were equal for all conditions compared to one

another.


5.3.1 DMF device fabrication and design

Digital microfluidics provides for the electrical manipulation of liquid,14

gas,131

plasma,132

and

sol and gel phase droplets59,99

(presented in chapter 4) across electrode array surfaces. For this

work we implemented a variation of a previously published device design42

with several notable

features: (1) the ground electrode of the top-plate composed of transparent indium tin oxide

97

permits imaging on-device, (2) interdigitated electrodes facilitate droplet movement during

actuation, (3) optical windows patterned on the bottom plate permit light microscopy imaging of

cells cultured on-device, and (4) the hydrophilic sites patterned on the device top-plate allow the

formation of microgels by passive dispensing (Figure 4-1).

5.3.2 On-demand cardiac microgel formation and assay

Two mechanisms of droplet splitting on DMF are described in the literature – active and passive

dispensing, the former relying on electrode actuation to split droplets,14

while the latter exploits

surface heterogeneities to generate sub-droplets by pinning through hydrophilic interactions (as

presented in chapter 2).42

In addition to being useful for high precision dispensing of reagents on

device, surface heterogeneities introduced by protein spotting,67

liftoff patterning,41,42

or

chemical functionalization63,80

have enabled applications including cell culture and DNA

immobilization. In chapter 4 of this thesis, we demonstrated biocompatibility of DMF and

passive dispensing for the formation of microgels on-demand (applied to a representative model

of epithelial morphogenesis). Here, we evaluated the compatibility of a similar method with

screening of functionality in 3D microtissues.

We selected a CM model system as there is great interest in evaluating the functional

characteristics of these cells, including contraction frequency and magnitude.133,134

Myocardial

tissue is organized in three-dimensions in vivo, but the vast majority of in vitro CM studies use

the conventional two-dimensional format.135

In the experiments reported here, we form 3D

microgels on-demand by passive dispensing sol-phase hydrogel material across hydrophilic sites,

98

Figure 5-1: Cardiac microgel formation on DMF. A sol-phase hydrogel droplet containing a

suspension of cardiac myocytes is drawn from a reservoir (A). The droplet is translated across

hydrophilic sites that are positioned (on the transparent top plate) above the optical windows on

the bottom plate (B-F). The white arrows indicate the direction of droplet movement. Scale bar =

4 mm.

99

followed by thermal cross-linking of the hydrogel (Figure 5-1). In this geometry, each microgel

is approximately 400 nL. When cells were seeded in varying densities, we observed two modes

of cellular arrangement throughout the construct. For lower initial seeding densities (up to 10 ×

106 cells/mL), cells were well dispersed throughout the 3D matrix (Figure 5-2A,B); however, at

higher initial seeding densities (20 × 106 cells/mL and above), cells formed aggregates within the

hydrogel matrix (Figure 5-2C).

At 3 or 4 days post-seeding, spontaneous contractions of 3D CM tissue formed from moderate-

density suspensions (10 × 106 cells/mL) were observed, followed by coordinated contractions on

day 5. Further, these constructs exhibited well connected 3D branched CM morphologies

throughout z (or vertical) axis of the microgel as observed by brightfield (Figure 5-3) and

confocal laser scanning microscopy (Figure 5-4, Figure 5-5). When seeded at lower densities (5

× 106 cells/mL) or higher densities (20 × 10

6 cells/mL), no or minimal spontaneous contractions

were observed. Thus, 3D CM constructs formed at 10 × 106 cells/mL were used for the

remainder of the experiments described here.

A functional response assay was performed to determine whether the cardiac microgel pillars

formed on demand were responsive to drug treatment. Specifically, cardiac microgels were

monitored by microscopy with contraction magnitude, duration, and frequency determined for

single cells within the construct (Figure 5-6). CMs evaluated in this manner exhibited a mean (±

1 S.D.) contraction magnitude of 2.4 µm ± 0.3 µm, a contraction duration of 0.11 s ± 0.02 s, and

a frequency of 50 beats per minute (BPM) ± 4.2 BPM. These data are consistent with literature

values for CM beating in vitro, which range from ~30 BPM for CMs cultured in 3D

100

Figure 5-2: Cell distributions in microgels as a function of seeding density. Phase contrast

images of micorgels formed with cells seeded at low (A), moderate (B), or high (C) density. The

cells are visible through windows on the bottom plate of the device. At low and modest densities,

cells are homogenously dispersed through the hydrogel. At high density, cell aggregation is

apparent. Scale bar = 250 µm.

Figure 5-3: Brightfield microscopy of CM microtissues. Images of CMs in a microgel formed

on-demand at 4x (A), 10x (B) and 20x (C) magnification. Images of cells in the same microgel at

20x magnification at z=40 (D), 120 (E), or 200 m (F).Scale bars = 50 µm.

101

Figure 5-4: Confocal images of cardiac microgel on DMF. A cardiac microgel was fixed and

stained (blue for nuclei and red for actin) after five days of culture on DMF. Imaging was

performed at 5× (A), 10× (B), and 20× (C) magnification, with elongated branched cells evident

at each level. Scale bars = 100 µm, 50 µm, and 25 µm respectively.

Figure 5-5: 3D cross-section of cardiac microgel. Cells were fixed and stained on DMF (blue

for nuclei, red for actin). Maximum combined exposure of an orthographic view in (A) z-x, (B)

x-y, and (C) y-z planes, demonstrates a well-connected network of CMs that are distributed in

3D space. Scale bar = 50 µm.

102

collagen/fibrin matrices130

to ~70-90 BPM for 2D cultured CMs.136,137

As cardiomyocytes are

known to be sensitive to the hormone and neurotransmitter epinephrine (EPI), we challenged

cardiac microgels with droplets of EPI and characterized their response: the magnitudes of

contraction, duration, and frequency increased by mean fold-values ± 1 S.D. of 1.9-fold ± 0.3,

1.8-fold ± 0.5, and 2-fold ± 0.2, respectively (Figure 5-6). These observations suggest that DMF

based cardiac microgel assays may be a useful new tool in understanding the functional

consequences of pharmacologic agents.

The microgel-on-demand method seems well-suited for parallel screening of conditions in 3D

cell constructs, as highlighted in Chapter 4. The new CM microgel-on-demand technique

described here joins a small group of other techniques directed to this purpose. For example,

Parker and colleagues122,126

recently described methods to form CM films that bend as a function

of the degree of CM contractions, a technique that should be readily adaptable to multiplexed

screening (by measuring bend frequency). This method is intriguing, but it is limited in that it is

not designed for 3D culture and it does not allow for each film to be addressed individually.

Likewise, Radisic and colleagues121

recently demonstrated a creative wire-based seeding strategy

to form functional cardiac ‘biowires’, but the protocol (as used now) is likely not amenable to

automation of high throughput tissue generation or individual ‘biowire’ treatment and analysis.

We posit that the CM microgel on-demand system presented here is unique in that it permits the

potential formation of multiple cardiac microtissues (e.g., 32 microgels, as in Figure 4-8A),

each of which can be individually addressed with independent treatments. Further, the footprint

of the microgels per unit area is equivalent to the conventional 96-well plate format, which

should facilitate translation to multiplexed screening. But unfortunately, the analysis

103

Figure 5-6: Cardiomyocyte function in microgels. Cardiac microtissues were cultured for four

days on DMF and evaluated before and after exposure to epinephrine. The areas of twenty

individual cells were calculated and plotted with respect to time – representative traces (blue-

unstimulated; red – EPI-stimulated) for one cell are shown in (A). Insets brightfield image shows

the contraction measurements performed on single cells. Black line indicates shape pre-

contraction, yellow line indicates shape post-contraction. These data were pooled and evaluated

for contraction frequency (B), contraction distance (C), and duration (D) for control and

epinephrine stimulated cells. Experiments performed in triplicate and error bars indicate ±1 S.D.

104

methods described here are too slow – new analytical tehcniques are needed to facilitate

efficient, multiplexed screening (as described below).

5.3.3 Higher efficiency analysis of cardiac microgel activity

The data in Figure 5-6 was collected using a standard single-cell image analysis technique that is

widely in CM research,138

but it requires excessive manual intervention for analysis. Specifically,

in our work, evaluating contractions on a cell-to-cell basis required ten minutes of analysis per

cell, which required >6 h to evaluate the two conditions represented in Figure 5-6. With

increasing numbers of conditions this method is prohibitive in terms of time required for

analysis. More importantly, the data is not representative of the tissue as a whole – evaluating the

activities of single cells misses the big-picture understanding of the functional behaviour of the

CM gel as a unit. For this reason we developed a faster method compatible with tissue-level

analysis to be combined with the novel DMF based cell handling technique. To this end, we

examined the ability to analyze CM tissue activity based on simple analysis of bright field

whole-image sequences. In an initial experiment we observed an initially beating cardiac

microgel during and after treatment with fixative. By calculating changes in pixel intensity

between frames, we generated an average intensity change for each image, which we termed the

cardiac activity coefficient (CAC). When CACs were plotted against time (Figure 5-7A) it was

clear that the morphological changes decreased, as is expected for the process of fixation.

Qualitative observation of image sequences demonstrated that with time, contraction frequency

across the microgel decreased until there was no longer any observable movement at 30 minutes

after treatment with fixative. This can be visualized as heat maps of activity corresponding to

105

time points at the beginning, middle, and end of the experiment (Figure 5-7B-D) that confirm

decreases and eventual cessation of contractile activity with exposure to fixative.

To test the CAC as a proxy for average CM contraction frequency, we applied it to the same data

used to generate the single-cell analysis shown in Figure 5-6. Images of unstimulated CM

microgels collected at 10× magnification had a mean (±R.S.D.) CAC of 1.44 ±.02, while

corresponding images of epinephrine-stimulated CM microgels had a mean (±R.S.D.) CAC of

1.58 ±.04 (Figure 5-8A). Each image evaluated the contributions of approximately 400-500 cells

(some perfectly in-focus, many slightly out of focus), and thus represented the behaviour of each

CM microgel as functional unit. The difference between the CACs for these two conditions was

statistically significant (p < 0.01). Heat maps representing activity generated from this data

demonstrate more regions of high activity in epinephrine treated cells than control (Figure

5-8B). Most importantly, this analysis was generated in a matter of seconds; to our knowledge,

this is the first report of such a tool. Moreover, this method allows for the evaluation of 3D tissue

activity as a whole. More work is required to validate and test the CAC method, but we propose

that this preliminary data is promising for the development of CAC as a tool for screening tissue-

level morphological changes.

5.3.4 Conclusion

Here, we demonstrated a DMF based microgel-on-demand method for high efficiency studies of

3D cardiac myocytes. This is the first microfluidic method that permits automated individual

addressability of microtissues for parallel and multiplexed experiments. Further, we developed a

new rapid analysis technique for evaluating the functional beading response of CMs grown in

106

3D. We expect that the implementation of DMF technology within high throughput screening

regimes will not only provide the benefit of reduced reagent consumption and improved

experimental workflow – but will enable physiologically relevant in vitro models of biology.

107

Figure 5-7: Cardiomyocyte activity coefficient. After five days of culture on DMF, a beating cardiac

microgel was treated with a fixative solution and observed at 10× magnification for 30 minutes (A). A heat

map was generated to represent cumulative cardiomyocyte activity coefficients at 1 (B), 15 (C), and 30 min

(D) after exposure to fixative. Heat maps are presented as overlays on phase contrast microscopic images.

Here blue indicates regions of low activity and red indicates high activity. With increased exposure to

fixative over time, reductions of activity are observed (as shown through reduced red regions in the heat

maps).

108

Figure 5-8: Cardiac activity coefficient of stimulated cells. CACs were calculated for control

and epinephrine-treated cardiac microgels, and the response (A) was significantly different by a

one-tailed t-test (p < 0.01). Heat maps representing cardiomyocyte activity generated for control

and epinephrine-stimulated microgels (B,C respectively), and brightfield images used for

analysis (D, E respectively). In these images red regions indicate higher activity, with blue

regions indicating lower activity. Scale bars = 50 µm.

109

Chapter 6. Concluding Remarks and

Perspectives on the Future

6.1 Conclusions

At the outset of carrying out the work described in this thesis, digital microfluidics as applied to

cell culture and analysis was very much in the “proof-of-principle” stage. Moreover, the

techniques being developed were constrained to specialized microfluidics laboratories such as

our own. Through the work presented here we have made significant contributions to (1)

improving DMF device functionality, (2) the development of DMF as a robust platform for 2D

cell culture, and (3) improving in vitro tools for the formation and analysis of 3D microtissues.

Chapter 2 describes the foundational development of hydrophilic site patterning within DMF

devices through the implementation of a fluorocarbon liftoff technique for improved device

functionality. This method overcomes limitations associated with inconsistent and incompatible

protein spotting techniques described in the initial reports of DMF based cell culture. We showed

this method was useful for the implementation of passive dispensing with high precision and

accuracy in both air and oil. The success of this technique is being appropriated in the Wheeler

Laboratory for a diverse array of applications ranging including impedance based cell

quantification139

, single cell analysis,140

immobilized immunoassays, janus nanoparticle

formation,141

and portable systems for in-the-field urinalysis by mass spectrometry.142

Further,

others have appropriated the method to study 2D constraints on cell morphology143

and for

increasing throughput of C. elegans droplet based behavioural assays.144

110

The method presented in Chapter 2 was fundamental to the development of robust DMF based

2D culture of primary cells, as described in Chapter 3. Previously published techniques required

protein spotting of extracellular matrix proteins; however, the previous techniques were

incompatible with these sensitive cell types. The implementation of the fluorocarbon liftoff

technique from Chapter 2 enabled the culture of primary cells and preliminary studies for the

culture of human embryonic stem cells in sub-microlitre droplets. Further we found that these

devices were amenable to integration with standard imaging equipment, permitting brightfield

microscopy and high resolution epifluorescent microscopy. This was the first platform to

integrate cell culture, fixing, and analysis completely on a single device. We demonstrated the

ability to run co-culture assays, furthering the versatility of DMF as a platform for cell biology.

Building on the ability to form discrete liquid structures on device (Chapter 2) combined with

the validation of biocompatibility with sensitive cell types (Chapter 3), we became interested in

exploring DMF as a platform for improved in vitro 3D cell culture models. In Chapter 4 we

present a novel technique termed ‘microgels-on-demand’ that for the first time permits the

formation of complex microscale hydrogel structures from thermally cross-linked hydrogel

materials. In this instance the unique properties of DMF enable a method that has not been

demonstrated with any other technique. With automation we demonstrate the ability to generate

high density individually addressable microgel structures in a wide range of geometries. Further,

the multiplexed liquid handling capabilities of DMF were used to demonstrate the formation of

combinatorial and composite microgels. Finally this system was used to recapitulate a model of

kidney epithelialization within a sub-microlitre droplet, and imaged on-device with confocal

microscopy.

111

The findings of Chapter 4 suggested to us that DMF microgels-on-demand could be used to

produce micro-scale functional tissues. In Chapter 5, we implemented this method for the

formation and assay of cardiac microgels. The tissues formed in this manner require minimal cell

material and presented spontaneous coordinated contractions after only a few days in culture.

Treatment with known accelerants demonstrated that these tissues were responsive and

representative of in vivo cardiomyocytes. Further we developeda new method for analyzing

whole tissue functional data.

6.2 Future perspectives

The specific contributions of this thesis build toward a broader vision of fully integrated micro-

scale systems for biological research. In parallel with other innovations in device fabrication and

automation, we are approaching the reality of autonomous bench top cell culture systems that are

accessible to the basic biologist. These technologies have to potential to disrupt the current state

of biological research workflow as they will enable improved consistency, by reducing manual

intervention, higher throughput, and lower-cost as a function of decreased reagent consumption.

We predict that the microgels-on-demand method presented in this thesis will enable the study of

3D microenvironment geometry influence on cell phenotype and behaviour, thereby improving

knowledge in tissue engineering and bridging the gap between in vitro and in vivo systems.

We are witnessing an ever increasing number of researchers joining the field of digital

microfluidics. Major and emerging players in the pharmaceutical, diagnostics, and electronics

industries are increasingly investing in the development of DMF. Life Technologies has invested

in cell culture applications, Abbot Diagnostics in immunoassays,96

and Sharp Laboratories in

thin film transitor based devices,16

Sofie Biosciences in radiotracer synthesis,98

and Advanced

112

Liquid Logic for dry blood spot analysis145

and PCR.146

The increasing interaction between

academic researchers and industrial players is driving rapid evolution in DMF technology and is

promoting its adoption within the research and clinical environments. With the development of

lower cost PCB15

and paper based devices147

combined with automation strategies, this

technology is well positioned to become accessible to a broad range of users in fields from

chemistry to biology.

With significant progress and interest over the past few decades, microfluidic technologies and

their application to biological questions are developing at an exponential rate. As those outside

the microfluidics community begin adopting these technologies we will find increasingly novel

biological questions to be answered at the micro scale, and find the adoption of these

technologies to have a broad impact at the level of basic research, drug discovery, and clinical

applications.

113

References

1. Schrödinger, E. What is life? The physical aspect of the living cell. (Cambridge

University Press, 1948).

2. Feynman, R. There is plenty of room at the bottom. Engineering and Science 23, 22–36

(1960).

3. Einstein, A. Investigations on the theory of brownian movement. (Dover Publications,

1956).

4. Berg, H. Random walks in biology. (Princeton University Press, 1993).

5. Taylor, G. Dispersion of soluble matter in solvent flowing slowly through a tube. Proc.

R. Soc. Lond. A Math. Phys. Sci. 219, 186–203 (1953).

6. Manz, A., Graber, N. & Widmer, H. M. Miniaturized total chemical analysis systems: A

novel concept for chemical sensing. Sensors and Actuators B: Chemical 1, 244–248 (1990).

7. Lemmo, A. V., Rose, D. J. & Tisone, T. C. Inkjet dispensing technology: applications in

drug discovery. Curr. Opin. Biotechnol. 9, 615–617 (1998).

8. Duffy, D. C., McDonald, J. C., Schueller, O. J. A. & Whitesides, G. M. Rapid

Prototyping of Microfluidic Systems in Poly(dimethylsiloxane). Analytical Chemistry 70, 4974–

4984 (1998).

9. Takatsy, G. The use of spiral loops in serological and virological micro-methods. Acta

Microbiol Acad Sci Hung 3, 191–202 (1955).

10. Squires, T. M. & Quake, S. R. Microfluidics: Fluid physics at the nanoliter scale. Rev.

Mod. Phys. 77, 977–1026 (2005).

114

11. Grimes, A. et al. Shrinky-Dink microfluidics: rapid generation of deep and rounded

patterns. Lab Chip 8, 170–172 (2007).

12. Hong, J., deMello, A. J. & Jayasinghe, S. N. Bio-electrospraying and droplet-based

microfluidics: control of cell numbers within living residues. Biomed Mater 5, 21001 (2010).

13. Choi, K. Digital Microfluidics.

14. Sung Kwon Cho, Hyejin Moon & Chang-Jin Kim. Creating, transporting, cutting, and

merging liquid droplets by electrowetting-based actuation for digital microfluidic circuits. J.

Microelectromech. Syst. 12, 70–80 (2003).

15. Jian Gong. Direct-Referencing Two-Dimensional-Array Digital Microfluidics Using

Multilayer Printed Circuit Board. Journal of Microelectromechanical Systems 17, 257–264

(2008).

16. Hadwen, B. et al. Programmable large area digital microfluidic array with integrated

droplet sensing for bioassays. Lab Chip 12, 3305–3313 (2012).

17. Pellat, H. Mesure de la force agissant sur les diélectriques liquides non électrisés placés

dans un champ électrique. C.R.Acad.Sci.Paris 119, 691–694

18. Lu, H.-W. et al. A Study of EWOD-Driven Droplets by PIV Investigation. Lab Chip 8,

456–461 (2008).

19. Poulos, J. L., Nelson, W. C., Jeon, T.-J., Kim, C.-J. ‘CJ’ & Schmidt, J. J. Electrowetting

on dielectric-based microfluidics for integrated lipid bilayer formation and measurement. Appl.

Phys. Lett. 95, 013706 (2009).

20. Wheeler, A. R., Moon, H., Kim, C.-J. ‘CJ’, Loo, J. A. & Garrell, R. L. Electrowetting-

Based Microfluidics for Analysis of Peptides and Proteins by Matrix-Assisted Laser

Desorption/Ionization Mass Spectrometry. Analytical Chemistry 76, 4833–4838 (2004).

115

21. Shah, G. J. EWOD-driven droplet microfluidic device integrated with optoelectronic

tweezers as an automated platform for cellular isolation and analysis. Lab on a Chip 9, 1732–

1739 (2009).

22. Song, J. H., Evans, R., Lin, Y.-Y., Hsu, B.-N. & Fair, R. B. A scaling model for

electrowetting-on-dielectric microfluidic actuators. Microfluid Nanofluid 7, 75–89 (2008).

23. Pollack, M. G., Fair, R. B. & Shenderov, A. D. Electrowetting-based actuation of liquid

droplets for microfluidic applications. Applied Physics Letters 77, 1725 (2000).

24. Paik, P., Pamula, V. K., Pollack, M. G. & Fair, R. B. Electrowetting-based droplet mixers

for microfluidic systemsElectronic supplementary information (ESI) available: six mpeg videos

showing some mixing schemes used in Fig. 7. See http://www.rsc.org/suppdata/lc/b2/b210825a/.

Lab Chip 3, 28 (2003).

25. Zhou, J. et al. Electrowetting-based multi-microfluidics array printing of high resolution

tissue construct with embedded cells and growth factors. Virtual & Phys. Prototyping 2, 217–223

(2007).

26. Bahadur, V. An energy-based model for electrowetting-induced droplet actuation.

Journal of Micromechanics and Microengineering 16, 1494–1503 (2006).

27. Verheijen, H. J. J. & Prins, M. W. J. Reversible Electrowetting and Trapping of Charge:

Model and Experiments. Langmuir 15, 6616–6620 (1999).

28. Chatterjee, D. Droplet-based microfluidics with nonaqueous solvents and solutions. Lab

on a Chip 6, 199–206 (2006).

29. Kang, K. H. How Electrostatic Fields Change Contact Angle in Electrowetting. Langmuir

18, 10318–10322 (2002).

116

30. Buehrle, J., Herminghaus, S. & Mugele, F. Interface Profiles near Three-Phase Contact

Lines in Electric Fields. Phys. Rev. Lett. 91, (2003).

31. Jones, T. B. An electromechanical interpretation of electrowetting. Journal of

Micromechanics and Microengineering 15, 1184–1187 (2005).

32. Abdelgawad, M., Park, P. & Wheeler, A. R. Optimization of device geometry in single-

plate digital microfluidics. J. Appl. Phys. 105, 094506 (2009).

33. Kang, K. H., Kang, I. S. & Lee, C. M. Wetting Tension Due to Coulombic Interaction in

Charge-Related Wetting Phenomena. Langmuir 19, 5407–5412 (2003).

34. Griffiths, D. J. Introduction to Electrodynamics. (Prentice-Hall, 1999).

35. Jones, T. B., Wang, K.-L. & Yao, D.-J. Frequency-Dependent Electromechanics of

Aqueous Liquids: Electrowetting and Dielectrophoresis. Langmuir 20, 2813–2818 (2004).

36. Choi, K., Ng, A. H. C., Fobel, R. & Wheeler, A. R. Digital microfluidics. Annu Rev Anal

Chem (Palo Alto Calif) 5, 413–440 (2012).

37. Walker, S. W., Shapiro, B. & Nochetto, R. H. Electrowetting with contact line pinning:

Computational modeling and comparisons with experiments. Phys. Fluids 21, 102103 (2009).

38. Sen, P. & Kim, C.-J. Capillary Spreading Dynamics of Electrowetted Sessile Droplets in

Air. Langmuir 25, 4302–4305 (2009).

39. Monroe, C. W., Urbakh, M. & Kornyshev, A. A. Double-Layer Effects in Electrowetting

with Two Conductive Liquids. J. Electrochem. Soc. 156, P21–P28 (2009).

40. Barbulovic-Nad, I., Yang, H., Park, P. S. & Wheeler, A. R. Digital microfluidics for cell-

based assays. Lab Chip 8, 519 (2008).

41. Srigunapalan, S., Eydelnant, I. A., Simmons, C. A. & Wheeler, A. R. A digital

microfluidic platform for primary cell culture and analysis. Lab on a Chip 12, 369 (2012).

117

42. Eydelnant, I. A., Uddayasankar, U., Li, B., Liao, M. W. & Wheeler, A. R. Virtual

microwells for digital microfluidic reagent dispensing and cell culture. Lab Chip 12, 750–757

(2012).

43. Au, S. H., Fobel, R., Desai, S. P., Voldman, J. & Wheeler, A. R. Cellular bias on the

microscale: probing the effects of digital microfluidic actuation on mammalian cell health,

fitness and phenotype. Integr. Biol. (2013). doi:10.1039/C3IB40104A

44. Pampaloni, F., Reynaud, E. G. & Stelzer, E. H. K. The third dimension bridges the gap

between cell culture and live tissue. Nature Reviews Molecular Cell Biology 8, 839–845 (2007).

45. Hall, H. G., Farson, D. A. & Bissell, M. J. Lumen formation by epithelial cell lines in

response to collagen overlay: a morphogenetic model in culture. Proceedings of the National

Academy of Sciences of the United States of America 79, 4672 (1982).

46. Töyli, M., Rosberg-Kulha, L., Capra, J., Vuoristo, J. & Eskelinen, S. Different responses

in transformation of MDCK cells in 2D and 3D culture by v-Src as revealed by microarray

techniques, RT-PCR and functional assays. Laboratory Investigation 90, 915–928 (2010).

47. Huh, D., Hamilton, G. A. & Ingber, D. E. From 3D cell culture to organs-on-chips.

Trends in Cell Biology 21, 745–754 (2011).

48. Unger, M. A., Chou, H.-P., Thorsen, T., Scherer, A. & Quake, S. R. Monolithic

Microfabricated Valves and Pumps by Multilayer Soft Lithography. Science 288, 113–116

(2000).

49. Tan, W. & Desai, T. A. Microfluidic Patterning of Cells in Extracellular Matrix

Biopolymers: Effects of Channel Size, Cell Type, and Matrix Composition on Pattern Integrity.

Tissue Engineering 9, 255–267 (2003).

118

50. Bauer, M., Su, G., Beebe, D. J. & Friedl, A. 3D microchannel co-culture: method and

biological validation. Integr. Biol. 2, 371 (2010).

51. Montanez-Sauri, S. I., Sung, K. E., Puccinelli, J. P., Pehlke, C. & Beebe, D. J.

Automation of three-dimensional cell culture in arrayed microfluidic devices. J Lab Autom 16,

171–185 (2011).

52. Tan, W.-H. & Takeuchi, S. Monodisperse Alginate Hydrogel Microbeads for Cell

Encapsulation. Advanced Materials 19, 2696–2701 (2007).

53. Tumarkin, E. et al. High-throughput combinatorial cell co-culture using microfluidics.

Integr Biol (Camb) 3, 653–662 (2011).

54. An, H. Z., Helgeson, M. E. & Doyle, P. S. Nanoemulsion Composite Microgels for

Orthogonal Encapsulation and Release. Advanced Materials 24, 3838–3844 (2012).

55. Jebrail, M. J. et al. A digital microfluidic method for dried blood spot analysis. Lab on a

Chip 11, 3218 (2011).

56. Shih, S. C. C. et al. Dried Blood Spot Analysis by Digital Microfluidics Coupled to

Nanoelectrospray Ionization Mass Spectrometry. Anal. Chem. 84, 3731–3738 (2012).

57. Yang, H., Mudrik, J. M., Jebrail, M. J. & Wheeler, A. R. A Digital Microfluidic Method

for in Situ Formation of Porous Polymer Monoliths with Application to Solid-Phase Extraction.

Anal. Chem. 83, 3824–3830 (2011).

58. Fiddes, L. K. et al. Hydrogel discs for digital microfluidics. Biomicrofluidics 6, 014112–

014112–11 (2012).

59. Luk, V. N., Fiddes, L. K., Luk, V. M., Kumacheva, E. & Wheeler, A. R. Digital

microfluidic hydrogel microreactors for proteomics. Proteomics 12, 1310–1318 (2012).

119

60. Abdelgawad, M., Freire, S. L. S., Yang, H. & Wheeler, A. R. All-terrain droplet

actuation. Lab Chip 8, 672–677 (2008).

61. Mousa, N. A. et al. Droplet-scale estrogen assays in breast tissue, blood, and serum. Sci

Transl Med 1, 1ra2 (2009).

62. Kirby, A. E. & Wheeler, A. R. Microfluidic origami: a new device format for in-line

reaction monitoring by nanoelectrospray ionization mass spectrometry. Lab Chip 13, 2533–2540

(2013).

63. Malic, L., Veres, T. & Tabrizian, M. Two-dimensional droplet-based surface plasmon

resonance imaging using electrowetting-on-dielectric microfluidics. Lab Chip 9, 473 (2009).

64. Kuehne, A. J. C. et al. A switchable digital microfluidic droplet dye-laser. Lab on a Chip

11, 3716 (2011).

65. Sia, S. K. & Whitesides, G. M. Microfluidic devices fabricated in Poly(dimethylsiloxane)

for biological studies. Electrophoresis 24, 3563–3576 (2003).

66. Delamarche, E., Bernard, A., Schmid, H., Michel, B. & Biebuyck, H. Patterned Delivery

of Immunoglobulins to Surfaces Using Microfluidic Networks. Science 276, 779 –781 (1997).

67. Barbulovic-Nad, I., Au, S. H. & Wheeler, A. R. A microfluidic platform for complete

mammalian cell culture. Lab Chip 10, 1536–1542 (2010).

68. Fouillet, Y. & Achard, J.-L. Microfluidique discrète et biotechnologie. Comptes Rendus

Physique 5, 577–588 (2004).

69. Chen, T., Su, C., Chih, H. & Yang, C. Selective wettability assisted nanoliter sample

generation via electrowetting-based transportation. ASME Conf. Proc. 147–153 (2007).

120

70. Vergauwe, N. et al. A versatile electrowetting-based digital microfluidic platform for

quantitative homogeneous and heterogeneous bio-assays. J. Micromech. Microeng. 21, 054026

(2011).

71. Witters, D. et al. Biofunctionalization of electrowetting-on-dielectric digital microfluidic

chips for miniaturized cell-based applications. Lab Chip 11, 2790–2794 (2011).

72. Bogojevic, D., Chamberlain, M. D., Barbulovic-Nad, I. & Wheeler, A. R. A digital

microfluidic method for multiplexed cell-based apoptosis assays. Lab on a Chip 12, 627 (2012).

73. Garyantes, T. Virtual wells for use in high throughput screening assays. (2003). at

<http://www.freepatentsonline.com/6565813.html>

74. Luk, V. N., Mo, G. C. & Wheeler, A. R. Pluronic Additives: A Solution to Sticky

Problems in Digital Microfluidics. Langmuir 24, 6382–6389 (2008).

75. Knovel Critical Tables. (Knovel, 2008).

76. Chang, Y.-H., Lee, G.-B., Huang, F.-C., Chen, Y.-Y. & Lin, J.-L. Integrated polymerase

chain reaction chips utilizing digital microfluidics. Biomed Microdevices 8, 215–225 (2006).

77. Miller, E. M. & Wheeler, A. R. A Digital Microfluidic Approach to Homogeneous

Enzyme Assays. Analytical Chemistry 80, 1614–1619 (2008).

78. Fair, R. B. et al. Chemical and biological applications of digital-microfluidic devices.

IEEE Design & Test of Computers 24, 10–24 (2007).

79. Wang, Y., Zhao, Y. & Cho, S. K. Efficient in-droplet separation of magnetic particles for

digital microfluidics. J. Micromech. Microeng. 17, 2148 (2007).

80. Malic, L., Veres, T. & Tabrizian, M. Biochip functionalization using electrowetting-on-

dielectric digital microfluidics for surface plasmon resonance imaging detection of DNA

hybridization. Biosensors and Bioelectronics 24, 2218–2224 (2009).

121

81. Fouillet, Y., Jary, D., Chabrol, C., Claustre, P. & Peponnet, C. Digital microfluidic design

and optimization of classic and new fluidic functions for lab on a chip systems. Microfluid

Nanofluid 4, 159–165 (2007).

82. Qiu, Q., Sayer, M., Kawaja, M., Shen, X. & Davies, J. E. Attachment, morphology, and

protein expression of rat marrow stromal cells cultured on charged substrate surfaces. J. Biomed.

Mater. Res. 42, 117–127 (1998).

83. Khademhosseini, A., Langer, R., Borenstein, J. & Vacanti, J. P. Microscale technologies

for tissue engineering and biology. PNAS 103, 2480–2487 (2006).

84. Finkel, N. H., Lou, X., Wang, C. & He, L. Barcoding the microworld. Anal. Chem. 76,

352A–359A (2004).

85. Lu, Y., Yin, Y. & Xia, Y. Three-Dimensional Photonic Crystals with Non-spherical

Colloids as Building Blocks. Advanced Materials 13, 415–420 (2001).

86. Balagaddé, F. K., You, L., Hansen, C. L., Arnold, F. H. & Quake, S. R. Long-term

monitoring of bacteria undergoing programmed population control in a microchemostat. Science

309, 137–140 (2005).

87. Liu, T. et al. A microfluidic device for characterizing the invasion of cancer cells in 3-D

matrix. Electrophoresis 30, 4285–4291 (2009).

88. Eddington, D. T. & Beebe, D. J. Flow control with hydrogels. Advanced Drug Delivery

Reviews 56, 199–210 (2004).

89. Agresti, J. J. et al. Ultrahigh-throughput screening in drop-based microfluidics for

directed evolution. Proceedings of the National Academy of Sciences 107, 4004–4009 (2010).

122

90. Leung, K. et al. A programmable droplet-based microfluidic device applied to

multiparameter analysis of single microbes and microbial communities. PNAS (2012).

doi:10.1073/pnas.1106752109

91. Edd, J. F. et al. Controlled encapsulation of single-cells into monodisperse picolitre

drops. Lab Chip 8, 1262–1264 (2008).

92. Dendukuri, D., Pregibon, D. C., Collins, J., Hatton, T. A. & Doyle, P. S. Continuous-flow

lithography for high-throughput microparticle synthesis. Nat Mater 5, 365–369 (2006).

93. Kloxin, A. M. et al. Responsive culture platform to examine the influence of

microenvironmental geometry on cell function in 3D. Integr Biol (Camb) 4, 1540–1549 (2012).

94. Nelson, W. C. et al. Incubated protein reduction and digestion on an electrowetting-on-

dielectric digital microfluidic chip for MALDI-MS. Anal. Chem. 82, 9932–9937 (2010).

95. Sista, R. S. et al. Heterogeneous Immunoassays Using Magnetic beads On a Digital

Microfluidic Platform. Lab Chip 8, 2188–2196 (2008).

96. Ng, A. H. C., Choi, K., Luoma, R. P., Robinson, J. M. & Wheeler, A. R. Digital

Microfluidic Magnetic Separation for Particle-Based Immunoassays. Analytical Chemistry 84,

8805–8812 (2012).

97. Jebrail, M. J. et al. Synchronized synthesis of peptide-based macrocycles by digital

microfluidics. Angew. Chem. Int. Ed. Engl. 49, 8625–8629 (2010).

98. Keng, P. Y. et al. Micro-chemical synthesis of molecular probes on an electronic

microfluidic device. PNAS 109, 690–695 (2012).

99. Fiddes, L. K. et al. Hydrogel discs for digital microfluidics. Biomicrofluidics 6, 14112–

1411211 (2012).

123

100. Axelrod, D., Koppel, D. E., Schlessinger, J., Elson, E. & Webb, W. W. Mobility

measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys J 16, 1055–

1069 (1976).

101. Zhang, Y. & Wang, T.-H. Full-Range Magnetic Manipulation of Droplets via Surface

Energy Traps Enables Complex Bioassays. Advanced Materials n/a–n/a (2013).

doi:10.1002/adma.201300383

102. Kleinman, H. K. & Martin, G. R. Matrigel: Basement membrane matrix with biological

activity. Seminars in Cancer Biology 15, 378–386 (2005).

103. Albrecht, D. R., Underhill, G. H., Wassermann, T. B., Sah, R. L. & Bhatia, S. N. Probing

the role of multicellular organization in three-dimensional microenvironments. Nat Meth 3, 369–

375 (2006).

104. Gel-forming structures and stages of red algal galactans of different sulfation levels -

Springer. at <http://link.springer.com/article/10.1007/s10811-007-9229-9/fulltext.html>

105. Park, J. H. et al. Microporous cell-laden hydrogels for engineered tissue constructs.

Biotechnology and Bioengineering 106, 138–148 (2010).

106. Rani, S. A., Pitts, B. & Stewart, P. S. Rapid Diffusion of Fluorescent Tracers into

Staphylococcus epidermidis Biofilms Visualized by Time Lapse Microscopy. Antimicrob Agents

Chemother 49, 728–732 (2005).

107. Kim, B. et al. Tuning Payload Delivery in Tumour Cylindroids using Gold Nanoparticles.

Nat Nanotechnol 5, 465–472 (2010).

108. Walker, G. M. et al. Effects of flow and diffusion on chemotaxis studies in a

microfabricated gradient generator. Lab Chip 5, 611–618 (2005).

124

109. Cornelissen, L. H., Bronneberg, D., Oomens, C. W. J. & Baaijens, F. P. T. Diffusion

measurements in epidermal tissues with fluorescent recovery after photobleaching. Skin Res

Technol 14, 462–467 (2008).

110. Tsang, V. L. et al. Fabrication of 3D hepatic tissues by additive photopatterning of

cellular hydrogels. FASEB J 21, 790–801 (2007).

111. Hui, E. E. & Bhatia, S. N. Micromechanical control of cell–cell interactions. PNAS 104,

5722–5726 (2007).

112. Ma, M. et al. Core–Shell Hydrogel Microcapsules for Improved Islets Encapsulation.

Advanced Healthcare Materials n/a–n/a (2012). doi:10.1002/adhm.201200341

113. Liu, T., Lin, B. & Qin, J. Carcinoma-associated fibroblasts promoted tumor spheroid

invasion on a microfluidic 3D co-culture device. Lab Chip 10, 1671–1677 (2010).

114. Elia, N. & Lippincott-Schwartz, J. Culturing Three Dimensional MDCK cells for

Analyzing Intracellular Dynamics. Curr Protoc Cell Biol Unit–4.22 (2009).

115. Au, S., Fobel, R. & Wheeler, A. Genome-level Cell Responses to Digital Microfluidic

Manipulation. Submitted

116. Matsunaga, Y. T., Morimoto, Y. & Takeuchi, S. Molding Cell Beads for Rapid

Construction of Macroscopic 3D Tissue Architecture. Advanced Materials 23, H90–H94 (2011).

117. Du, Y., Lo, E., Vidula, M. K., Khabiry, M. & Khademhosseini, A. Method of Bottom-Up

Directed Assembly of Cell-Laden Microgels. Cell Mol Bioeng 1, 157–162 (2008).

118. Gilbert, P. Personal communication.

119. Zaman, M. H. et al. Migration of tumor cells in 3D matrices is governed by matrix

stiffness along with cell-matrix adhesion and proteolysis. PNAS 103, 10889–10894 (2006).

125

120. Harary, I. & Farley, B. In vitro studies on single beating rat heart cells: I. Growth and

organization. Experimental Cell Research 29, 451–465 (1963).

121. Nunes, S. S. et al. Biowire: a platform for maturation of human pluripotent stem cell-

derived cardiomyocytes. Nat Meth advance online publication, (2013).

122. Agarwal, A., Goss, J. A., Cho, A., McCain, M. L. & Parker, K. K. Microfluidic heart on a

chip for higher throughput pharmacological studies. Lab Chip (2013). doi:10.1039/C3LC50350J

123. Köster, S. et al. Drop-based microfluidic devices for encapsulation of single cells. Lab

Chip 8, 1110–1115 (2008).

124. Khademhosseini, A. et al. Microfluidic patterning for fabrication of contractile cardiac

organoids. Biomed Microdevices 9, 149–157 (2007).

125. Khanal, G., Chung, K., Solis-Wever, X., Johnson, B. & Pappas, D. Ischemia/reperfusion

injury of primary porcine cardiomyocytes in a low-shear microfluidic culture and analysis

device. Analyst 136, 3519–3526 (2011).

126. Grosberg, A., Alford, P. W., McCain, M. L. & Parker, K. K. Ensembles of engineered

cardiac tissues for physiological and pharmacological study: Heart on a chip. Lab Chip 11,

4165–4173 (2011).

127. Annabi, N. et al. Hydrogel-coated microfluidic channels for cardiomyocyte culture. Lab

Chip (2013). doi:10.1039/C3LC50252J

128. Kloß, D. et al. Microcavity array (MCA)-based biosensor chip for functional drug

screening of 3D tissue models. Biosensors and Bioelectronics 23, 1473–1480 (2008).

129. Fobel, R., Fobel, C. & Wheeler, A. R. DropBot: An open-source digital microfluidic

control system with precise control of electrostatic driving force and instantaneous drop velocity

measurement. Applied Physics Letters 102, 193513–193513–5 (2013).

126

130. Boudou, T. et al. A Microfabricated Platform to Measure and Manipulate the Mechanics

of Engineered Cardiac Microtissues. Tissue Engineering Part A 18, 910–919 (2012).

131. Zhao, Y. & Cho, S. K. Micro air bubble manipulation by electrowetting on dielectric

(EWOD): transporting, splitting, merging and eliminating of bubbles. Lab Chip 7, 273–280

(2007).

132. Fan, S.-K. et al. Atmospheric-pressure microplasma in dielectrophoresis-driven bubbles

for optical emission spectroscopy. Lab on a Chip 12, 3694 (2012).

133. Hansen, A. et al. Development of a Drug Screening Platform Based on Engineered Heart

Tissue. Circulation Research 107, 35–44 (2010).

134. Iyer, R. K., Chiu, L. L., Reis, L. A. & Radisic, M. Engineered cardiac tissues. Current

Opinion in Biotechnology 22, 706–714 (2011).

135. Zimmermann, W. H. et al. Three-dimensional engineered heart tissue from neonatal rat

cardiac myocytes. Biotechnology and Bioengineering 68, 106–114 (2000).

136. Fukuhara, S. et al. Direct cell-cell interaction of cardiomyocytes is key for bone marrow

stromal cells to go into cardiac lineage in vitro. J Thorac Cardiovasc Surg 125, 1470–1480

(2003).

137. Webster, D. R. & Patrick, D. L. Beating rate of isolated neonatal cardiomyocytes is

regulated by the stable microtubule subset. Am J Physiol Heart Circ Physiol 278, H1653–H1661

(2000).

138. Hersch, N. et al. The constant beat: cardiomyocytes adapt their forces by equal

contraction upon environmental stiffening. Biology Open (2013). doi:10.1242/bio.20133830

127

139. Shih, S. C. C., Barbulovic-Nad, I., Yang, X., Fobel, R. & Wheeler, A. R. Digital

microfluidics with impedance sensing for integrated cell culture andanalysis. Biosensors and

Bioelectronics 42, 314–320 (2013).

140. Ng, A. H. & Chamberlain, D. Personal communication.

141. Ng, A. & Sykes, E. Personal communication.

142. Lafrenière, N. M. & Kirby, A. E. Personal communication.

143. Pelletier, L. Personal communication.

144. Palanski, K. & Ryu, W. Personal communication.

145. Sista, R. S. et al. Multiplex newborn screening for Pompe, Fabry, Hunter, Gaucher, and

Hurler diseases using a digital microfluidic platform. Clin. Chim. Acta 424C, 12–18 (2013).

146. Schell, W. A. et al. Evaluation of a digital microfluidic real-time PCR platform to detect

DNA of Candida albicans in blood. Eur. J. Clin. Microbiol. Infect. Dis. 31, 2237–2245 (2012).

147. Fobel, R. Personal communication.

128

APPENDIX A

Figure A.1: Spheroid identification after manipulation. For deformation studies

individual spheroids were imaged before manipulation by ALHR or DMF by light

microscopy (indicated with arrows). To identify spheroids post-manipulation the

automated position coordinates on the automated microscope were used to find the x-y

coordinates. Then local fiduciary markers (particularly other spheroid structures,

indicated by asterisks in the figure) were used to determine exact spheroid location.

Digital Microfluidics for ... - University of Toronto

Documents

Transcript of Digital Microfluidics for ... - University of Toronto