Microfluidic Device for Co-culturing Lung Airway Cells on a ......ii Microfluidic Device for Co-culturing Lung Airway Cells on a Suspended Hydrogel: Towards a Biomimetic Lung Airway

Microfluidic Device for Co-culturing Lung Airway Cells

on a Suspended Hydrogel: Towards a Biomimetic

Lung Airway Model

by

Mouhita Humayun

A thesis submitted in conformity with the requirements for the degree of Master of Applied Science

Department of Mechanical & Industrial Engineering University of Toronto

© Copyright by Mouhita Humayun (2016)

ii

Microfluidic Device for Co-culturing Lung Airway Cells

on a Suspended Hydrogel: Towards a Biomimetic

Lung Airway Model

Mouhita Humayun

Master of Applied Science

Department of Mechanical Engineering University of Toronto

2016

Organ-on-a-chip technologies have made significant strides in recapitulating the complex

multicellular 3D architectures, tissue-tissue interfaces and the physiologically relevant

mechanical and biological properties of in vivo tissue microenvironments. To accelerate current

drug development processes, new and improved in vitro models that reconstitute the complex in

vivo microenvironment of human tissues are needed. Local disruptions to the lung airway

epithelium are known to elicit specific cellular responses leading to the progression of a myriad

of respiratory diseases, making it a crucial target for conducting disease mechanism and drug

development studies. We aimed to develop and perform preliminary assessment on a

microfluidic-based in vitro model that mimics the in vivo tissue microenvironments of the human

airway. By leveraging microfabrication techniques and surface tension principles to integrate

co-culture of airway cells on stable air- liquid interfaces, we demonstrated a microfluidic platform

that allows more efficient 3D culture experiments and enables high-throughput co-culture

studies of the airway tissue.

iii

To Mom and Dad,

for picking me up every time I fall

and for always believing in me.

iv

Acknowledgements

The past two years have been a tremendous learning experience for me and it is thanks in large

part to the special people who supported, challenged and stuck with me along the way. First, I

would like to express my deepest appreciation to my supervisor, Dr. Edmond Young, for

providing me with the opportunity to work in the IBMT lab and for giving me a chance to work on

this amazing project. His intellectual guidance and support made it possible for me to continue

this research even at points where moving forward seemed impossible to me. I am also grateful

for his patience and continuous emotional support during times of struggle and hardship. Thank

you, also, to my committee members, Prof. Chung-Wai Chow and Prof. Lidan You, for making

time for me and for providing valuable feedback on my thesis.

I would like to extend my sincerest gratitude to all the graduate students and post-docs that I

have shared our lab with during this project: Dr. Alwin Wan, Dr. Deepika Devadas, Thomas

Moore, Tobe Madu, Daniel Konstantinou and Noosheen Walji. Aside from all the help they have

provided me with by teaching me protocols and lending a helping hand during experiments, they

have made my experience in the lab incredibly enjoyable. Alwin- thank you for your advice on

microfabrication techniques and graduate school in general. Without your support, this project

would not be where it is today. Deepika- thank you for allowing me to bounce ideas off of you

and for putting up with my endless questions. Tom – thank you for taking the time to listen to me,

for challenging my ideas and for helping me bring out the core of any strong argument. I’ve

learned a lot about how to conduct “good research” from our conversations. Tobe – I’ve really

enjoyed our long strings of back-and-forth dialogue about the craziest things. Our conversations

have inspired me to pay attention to and appreciate qualities in people that I would normally

neglect. Daniel – thanks for always brightening up my day with your awesome sense of humour

and positive attitude. Noosh- thanks for always listening to me and supporting me, even when

v

my ideas were “unconventional”. I would also like to thank Danielle Machado, who helped me

with data collection, during her time in the lab as an undergraduate research student. It was a

pleasure to have worked alongside all of my labmates and I appreciate all the great

conversations I have had with them.

I owe a great debt of gratitude to Dr. Lindsey Fiddes and Dr. Dan Voicu who went out of their

way countless times to help me with my imaging issues. I thank them both for donating their time

to me and for their insights into my project.

I would like to thank my best friends Rahanuma Wafa and Priya Patel for making my time

outside of school so joyous and for always keeping my spirits up. I thank them both for never

judging, for always being honest and for never failing to provide much-needed emotional

support.

Finally, I would very much like to thank my family. My Mom, Dad and my sister have always

supporting me in everyway I can imagine. To Muna- thanks for always picking up my calls. To

Mom and Dad- thank you for all of your love and support. Thank you for teaching me to always

believe in myself, to persevere and to shoot for the moon.

vi

Table of Contents

1.0 Introduction ......................................................................................................................... 1

2.0 Background - Literature Review .......................................................................................... 3

2. 1 Anatomy of the Respiratory System ................................................................................ 4

2.2 Basic Morphology of Airway Tissues in the Tracheobronchial Tree ................................. 6

2.2.1 Basic Tissue Structures ............................................................................................. 7

2.2.2 Tissue Morphology of the Trachea ............................................................................ 8

2.2.3 Tissue Morphology of the Bronchi and Bronchioles ................................................... 8

2.3 Pathophysiology .............................................................................................................. 9

2.3.1 Airway Defense Mechanisms .................................................................................. 10

2.3.2 Asthma Pathophysiology ......................................................................................... 12

2. 4 Lung Tissue Models: In Vivo vs. In Vitro ....................................................................... 13

2.4.1 Selection of Cell Types and Cell sources ................................................................ 16

2.4.2 Selection of Physiologically Relevant Scaffold ........................................................ 18

2.4.3 Microfluidic Lung Models (Organ-on-a-chip systems) .............................................. 19

2.5 Thesis Objectives .......................................................................................................... 26

2.6 Future Outlook ............................................................................................................... 28

3.0 Airway-on-a-Chip Design and Fabrication ......................................................................... 29

3.1 Microfluidic Design ........................................................................................................ 31

3.2 Device Microfabrication and Assembly .......................................................................... 31

3.3 Materials and Methods .................................................................................................. 33

3.3.1 Device Design and Fabrication ................................................................................ 33

3.3.2 Solvent-Assisted Thermal Bonding ......................................................................... 35

3.4. Results ......................................................................................................................... 36

4.0 Suspended Microfluidic Principles ..................................................................................... 37


4.1.1 Device Design ......................................................................................................... 42

4.1.2 Micromilling ............................................................................................................. 42

4.1.3 Plasma Treatment ................................................................................................... 42

4.1.4 Preparation of Solutions .......................................................................................... 43

4.1.5 Pressure-Driven Flow Experiment ........................................................................... 43

4.3 Results and Discussion ................................................................................................. 43

4.3.1 Contact Angle ......................................................................................................... 43

4.3.2 Pressure-Driven Flow Experimental Data for 25% IPA Solution and DI Water ........ 44

4.3.3 Comparison between Theoretical and Experimental Pressure-Driven Flow Data for

25% IPA Solution and DI Water ....................................................................................... 48

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5.0 Preliminary Assessment of the Lung Airway Device .......................................................... 51


5.1.1 Device Design and Fabrication ................................................................................ 55

5.1.2 Cells and Cell Culture.............................................................................................. 55

5.1.3 Experimental Preparation ........................................................................................ 56

5.1.4 Cell Adhesion on Hydrogel Study ............................................................................ 57

5.1.5 Long-term Cell Culture in Microfluidic Device .......................................................... 60

5.1.6 Cell Seeding Density Study ..................................................................................... 61

5.1.7 Calu-3 epithelium barrier structure .......................................................................... 62

5.1.8 Actin labelling in HBSMCs ....................................................................................... 63

5.1.9 Co-culturing Calu-3 cells and HBSMCs ................................................................... 63

5.1.10 Immunostaining Co-culture Samples ..................................................................... 64

5.1.11 Microscopy of Immunolabeled Samples and Data Analysis ................................... 64

5.2 Results .......................................................................................................................... 67

5.2.1 Cell Adhesion Study ................................................................................................ 67

5.2.2 Calu-3 Monolayer Formation and seeding density ................................................... 76

5.2.3 HBSMC Seeding Density Study .............................................................................. 78

5.2.4 Monoculture Characterization.................................................................................. 79

5.2.5 Co-culture Characterization ..................................................................................... 84

5.3 Discussion ..................................................................................................................... 90

5.3.1 Calu-3 Cell Adhesion .............................................................................................. 90

5.3.2 HBSMC Cell Adhesion ............................................................................................ 92

5.3.3 Cell Seeding Density Study ..................................................................................... 93

5.3.4 Monoculture and Co-culture Characterization ......................................................... 94

6.0 Conclusions and Recommendations ................................................................................. 97

6.1 Summary ....................................................................................................................... 97

6.2 Recommendations ......................................................................................................... 99

7.0 Bibliography .................................................................................................................... 101

Appendix A ........................................................................................................................... 107

A1.0 Theoretical Approach to Characterizing Suspended Microflow with Applied Pressure 107

A1.1 Free Energy Calculation ........................................................................................ 107

A1.2 Analytical Model of Pressure Driven Suspended Microflow .................................... 109

A2.0 Dynamic Approach of Pressure Driven Suspended Microflow between Parallel Plates

.......................................................................................................................................... 110

A2.1 Description of the Physical Problem....................................................................... 110

A2.2 The General Governing Equation of Fluid Motion .................................................. 111

A2.3 Pressure, Capillary and Drag Force Calculation .................................................... 111

A2.4 Force Balance ....................................................................................................... 111

A2.5 Force Balance Neglecting the Inertial force ............................................................ 112

A2. 6 Velocity Profile ...................................................................................................... 112

A3.0 Flowrate to Pressure drop ......................................................................................... 113

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A3.1 Relationship Between Pressure Drop and Flowrate in a Microfluidic Channel ........... 113

A3.2 Volumetric Flowrate Calculation from Velocity Profile ................................................ 113

A3.3 Flowrate Simplification ........................................................................................... 114

A4.0 Frequency distribution of HBSMC alignment angle ................................................... 117

A5.0 Device fabrication and Cell Culture Protocol ............................................................. 120

A5.1 Device Fabrication Protocol ................................................................................... 120

A5.2 Cell Culture Protocol .............................................................................................. 121

A6.0 Cell Adhesion Data.................................................................................................... 126

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List of Tables

Table 4.1 Physical properties of 25% IPA solution and DI water required for theoretical

solutions……………………………………………………………………………...44

Table 4.2 Experimental data for 25% IPA solution and wate………………………………..45

Table 5. 1 Compositions ECM hydrogels tested for cell adhesion study. …………….……58

x

List of Figures

Figure 2.1 The anatomy of the upper and lower respiratory tract............................................... 5

Figure 2.2 Illustration of various cellular shape and organization of epithelial cells.................... 7

Figure 2.3 Basic histology of airway tissues. ............................................................................. 9

Figure 2 .4 Microfluidic in vitro airway models. .........................................................................25

Figure 2.5 Simplified model of the airway wall tissue structure.. ...............................................27

Figure 3.1 PMMA microfluidic device for co-culturing airway cells. ...........................................32

Figure 3.2 Microfabrication of PMMA microfluidic device for co-culturing airway cells. .............33

Figure 3.3 Schematic of device design and parameters for a 6 system PMMA co-culture

device . ...................................................................................................................35

Figure 3.4 Photograph of assembled PMMA microfluidic device. .............................................36

Figure 4.1 Schematic of a simplified bronchiole wall (left) with a ciliated monolayer epithelium

surrounded by a thin layer of extracellular matrix wrapped around by a band of

smooth muscle tissue. ............................................................................................38

Figure 4.2 Schematic of pressure driven suspended microflow between parallel walls for

channel width/height aspect- ratio greater than 1. ...................................................41

Figure 4.3 Photographs of contact angles formed by DI water (blue) and 25% IPA in DI water

(red) on a PMMA surface, before (A) and after (B) plasma treatment. ....................44

Figure 4.4 Individual frames from real-time video capture of pressure-driven suspended

microflow between parallel walls for 25% IPA in DI water. ......................................46

Figure 4.5 Individual frames from real-time video capture of pressure-driven suspended

microflow between parallel walls for water. .............................................................47

Figure 4.6 Penetration distance of the liquid front versus time for 25% IPA (orange curve), and

water (blue curve) with a contact angle of 25.3o and 4.8o (after plasma- treatment)

respectively on the wall surface. .............................................................................48

Figure 4.7 Velocity of 25% IPA (orange curve), and water (blue curve) versus time, with a

contact angle of 25.3o and 4.8o respectively on the wall surface. ............................49

Figure 5.1 Two layer PMMA microfluidic device for cell adhesion study. ..................................56

Figure 5.2 Calu-3 cells cultured in a T-25 flask form networked islands. Scale bar = 400µm. ...60

Figure 5.3 Sample preparation for microscopy of immunolabeling experiments. ......................66

Figure 5.4 Immunostaining of HBSMCs culture on the underside of an ECM hydrogel

suspended in a microfluidic device for 7 days. ........................................................67

Figure 5.5 Calu-3 epithelial cell adhesion on ECM hydrogel in a PMMA microfluidic device. ....70

Figure 5.6 Area covered by adhered Calu-3 cells (%) on ECM hydrogel, containing various

compositions of protein, inside a PMMA airway microfluidic device. .......................71

Figure 5.7 Comparison of area covered by adhered Calu-3 cells (%) on various ECM hydrogel

compositions between day 2 and 7. ........................................................................71

Figure 5.8 Fluorescence image of HBSMCs on different hydrogel compositions cultured

inside a PMMA microfluidic device for 7 days. ........................................................73

Figure 5.9 Fluorescence image of HBSMCs on different hydrogel compositions cultured

inside a PMMA microfluidic device for 2 and 7 days. ...............................................74

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Figure 5.10 Number of adhered HBSMCs on various ECM hydrogels during 7-day culture

period......................................................................................................................75

Figure 5.11 Cell Adhesion and growth of HBSMCs cultured for 7 days on various ECM

hydrogels. ...............................................................................................................75

Figure 5.12 Area of ECM hydrogel covered by cultured Calu-3 epithelial inside a PMMA

microfluidic device...................................................................................................77

Figure 5.13 Area covered by adhered Calu-3 cells (%) on a suspended ECM hydrogel, inside

a PMMA airway microfluidic device, cultured over a 21-day period. ........................78

Figure 5.14 Immunostaining of Calu-3 and HBSMCs co-cultured on opposing sides of an

ECM hydrogel suspended in a microfluidic device. .................................................79

Figure 5.15 Immunofluorescence staining of Calu-3 monolayers, cultured on a suspended

ECM hydrogel inside a microfluidic device, at day 7, 14 and 21. .............................82

Figure 5.16 Immunostaining of HBSMC monocultures on the underside of a suspended

hydrogel in a microfluidic device, at day 4, 7 and 14. ..............................................83

Figure 5.17 Immunolabelled fluorescent images of HBSMC monoculture at day 14, culture on

the underside of a suspended hydrogel in a microfluidic device. .............................85

Figure 5.18 Polar graphs showing frequency distribution of the angle of HBSMC alignment

with respect to the inlet-outlet axis of the microfluidic channel. ...............................85



Figure 5.20 Polar graphs showing frequency distribution of the angle of HBSMC alignment

with respect to the inlet-outlet axis of the microfluidic channel. ...............................88



xii

List of Abbreviations

AC acellular AECs alveolar epithelial cells ALI air-liquid interface AM alveolar macrophages ASM airway smooth muscle BSC biological safety cabinet CAD computer-aided design CNC computer numerical control COC cyclin olefin copolymer COPD chronic obstructive pulmonary disease DI deionized ECM extracellular matrix FN fibronectin HBSMCs human bronchial smooth muscle cells HBTE human bronchial tracheobronchial epithelial hESCs human embryonic stem cells HPMECs human pulmonary microvascular endothelial cells ICAM-1 intercellular adhesion molecule 1 IL-2 interleukin-2 IPA isopropyl alcohol iPSCs induced pluripotent stem cells PBS phosphate buffered saline PDMS polydimethylsiloxane PFA paraformaldehyde PGA poly(glycolic) acid PLA poly (lactic-co-glycolic) acid PMMA poly (methyl methacrylate) PS polystyrene SCF spontaneous capillary flow TEER transepithelial electrical resistance TJs tight junctions VCAM-1 vascular adhesion molecule 1 α-SMA α-smooth muscle actin

1

1.0 Introduction

Worldwide, more than 1 billion people suffer from chronic respiratory diseases and each year 4

million lives are lost prematurely from these diseases (Yorgancioglu et al., 2016). Chronic

respiratory diseases affect the airways of the lung, with smoking, pollution and genetic factors

being important risk factors that increase an individual’s chance of asthma or cystic fibrosis.

From a treatment perspective, current preclinical drug development processes, in developed

countries rely on expensive and time consuming animal testing that often fail to accurately

predict human responses. Majority of higher failure rates at clinical stages of drug development

are due to a lack of drug safety and efficacy (Arrowsmith and Miller 2013) Changes in

development strategies, particularly intervention in the research and development stages, are

required to avoid these costly failures. To improve prediction rates and reliability of drug safety

and efficacy in preclinical studies, new technologies that enable the study of complex human

physiology are needed.

Advances in the areas of microengineered technologies and microfluidic systems have enabled

us to artificially engineer a more realistic cell culture microenvironment by building miniaturized

perfusion bioreactors with continuous supply of cell nutrients and oxygen as well as by

integrating technologies that provide cells with various physiological stimuli. In addition, these

technologies can be leveraged to build organ-level tissue structures and geometries with the

potential for parallelization and increased throughput. Consequently, organ-on-a-chip and

1.0 Introduction 2

body-on-a-chip systems have emerged as competing platforms to animal models and

conventional 2D cultures for studying various in vivo processes including tissue functionality as

wells as responses to drugs and environmental toxins. Noteworthy examples of organ-on-a-chip

systems developed for conducting lung-related studies include the reconstituted

alveolar-capillary interface (Huh et al., 2010) and the small airway-on-a-chip (Benam et al.,

2010), both of which incorporate an extracellular matrix coated porous substrate separating the

culture of two different cell types to recapitulate the tissue-tissue interface of the in vivo

microenvironment. These biomimetic microdevices have demonstrated intriguing organ-level

physiological functions, including inflammatory responses to pathogens, epithelial and

endothelial barrier integrity and provided insights into the mechanisms of human inflammatory

lung disorders. Despite such exciting developments in these recapitulations, challenges still

remain in making these technologies amenable to widespread usage in the laboratory and for

applications in preclinical stages of drug development.

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2.0 Background - Literature Review

The purpose of this chapter is to outline the basic principles behind current approaches used to

better understand common chronic respiratory diseases that affect the airways and identify

challenges in the area of in vitro modelling that have yet to be addressed. We look at emerging

and conventional in vitro technologies that study the mechanisms of common respiratory

responses and compare their advantages and disadvantages.

To better equip us for assessing these technologies, we review the anatomy and basic functions

of the respiratory system, with a focus on the tracheobronchial tree of the lower respiratory

system. This is followed by a description of relevant tissue morphologies and cellular

components to enhance our understanding of the role of healthy airway tissues. We then look at

common airway defense mechanisms and their involvement in the progression of common

chronic respiratory diseases worldwide such as asthma and chronic obstructive pulmonary

disease to identify relevant components in modelling the airway and its diseases. Lastly, we

review current in vitro technologies, more specifically organ-on-a-chip systems, to reflect on

current progress and discuss potential technological advances. In the end, we propose an in

vitro system that incorporates relevant structures of the airway and addresses some of the

limitations of existing technologies.

2.0 Background - Literature Review 4

2. 1 Anatomy of the Respiratory System

The respiratory system is made up of organs and structures that enable the exchange of carbon

dioxide and oxygen between the circulating blood and the atmosphere at a rate rapid enough to

accommodate the body's needs. As shown in Figure 2.1, the air enters the body through the

nose or mouth and passes through the larynx, commonly known as the voice box, then down

the trachea before splitting into the bronchial tubes and eventually reaching the gas exchange

region of the lungs.

The main structures of the respiratory system can be divided into two groups based on their

functions. The first group is called the conducting zone, which serves the function of carrying the

inhaled air to and from the site of gas exchange while filtering and regulating the temperature

and humidity of the inhaled air (Lechner, Matuschak, and Brink 2011). Inclusive in the

conducting zone are airway structures, such as the nasal cavity, larynx, trachea, bronchi,

bronchioles and terminal bronchioles. The structures from the nasal cavity to the larynx is

primarily involved in humidifying and warming the air while the tracheobronchial tree

predominately contributes to the gas transfer and filtration tasks.

The second zone is called the respiratory parenchyma, which includes the final divisions of the

terminal bronchioles called respiratory bronchioles, alveolar ducts and alveoli. These structures

are primarily a collection of thin membrane walls that support molecular diffusion between the

inhaled air and blood flowing through the underlying vasculature (Rhoades and Bell 2009).

The tracheobronchial tree is the functional component of the respiratory system that allows

transport of air from the upper respiratory tract to the lung parenchyma, and includes structures

from the trachea to the alveoli. The trachea, commonly referred to as the windpipe, is the largest

airway and has several evenly spaced cartilage rings that keep the windpipe open for breathing

(Rogers 2011). The bronchi are two airway tubes that stem off from the trachea into


progressively smaller airways, called bronchioles, inside the lung in a tree-like pattern.

Together, the bronchi and the bronchioles ensure that the partially filtered air from the trachea is

delivered to all regions of the lung and further refined by the mucus-producing ciliated epithelium

that line the inner walls of these airways. The outer walls of the bronchi and bronchioles are

wrapped with muscle tissue which help control the flow of air travelling to the gas exchange

region (Rhoades and Bell 2009).

The bronchiole tubes lead to small air sacs called alveoli, which have very thin walls that allow

for gas exchange to occur in the lungs. The alveoli are surrounded by tiny blood vessels called

capillaries that are connected to a larger network of blood vessels called pulmonary arteries and

veins (Rogers, 2011). The pulmonary arteries carry carbon dioxide-rich blood from the body to

the capillaries that surround the alveoli where the oxygen from the air moves into the blood in

the capillaries and the carbon dioxide moves into the air. The oxygen-rich blood is then

delivered to the body through the heart where it is needed for cellular respiration. In this

Figure 2. 1 The anatomy of the upper and lower respiratory tract. Source: (Molnar and Gair 2015).


process, cells use the available oxygen to break down nutrients, which releases energy for

sustaining life while producing carbon dioxide.

Other structures of the respiratory system, commonly referred to as the muscles of respiration,

including the diaphragm and the intercostal muscles surrounding the lung lobes, help to expand

and compress the lungs during inhalation and exhalation (Rhoades and Bell 2009).

2.2 Basic Morphology of Airway Tissues in the Tracheobronchial Tree

Daily inhalation amounts to thousands of liters of ambient air passing through the airways

containing various types of microbes, harmful particulates, and noxious gases. Remarkably

enough, for most individuals the functional tissues of the lung, involved in gas transfer and

exchange, remain intact and unaffected unless the airway defence mechanisms are

compromised (Lechner, Matuschak, and Brink 2011). The airway defence mechanisms include

both physical and biological processes, discussed later in this section.

The basic morphology at each level of the tracheobronchial tree is similar, starting with a surface

epithelium lining the inner airway lumens, primarily consisting of ciliated and mucus-secreting

cells that overlay sub-epithelial tissue largely composed of glands and connective tissues

(Plopper and Hyde 2015). The cellular composition and tissue content vary at different levels of

the tracheobronchial tree.

Before diving into the specific tissue morphologies of the airway structures under consideration,

it is important to briefly review relevant animal tissue types and structures to ease understanding

of airway morphology.


2.2.1 Basic Tissue Structures

In general, animal tissues are classified into four types: epithelial, connective, muscular and

nervous. Although some airway tissues may include all four of these tissue types, much of the

airway is predominately composed of epithelial and connective tissue.

Epithelial tissue forms the inner lining of the airways in the upper and lower respiratory tract.

These tissues are tightly packed where one surface of the epithelium is always exposed to open

space while the opposing side faces an extracellular fibrous basement membrane that

separates the underlying tissues. These tissues are the airway's first line of defence against

physical, chemical, and biological disturbances. Epithelial cells primarily take on three principal

shapes: squamous, columnar and cuboidal (Figure 2.2). Each of these shapes can be arranged

in a single or multiple layers, classified as simple and stratified epithelium respectively.

In the context of airways, connective tissues generally support or separate other tissue layers of

the airways. The main components of connective tissues are various cells, cartilage, elastic and

collagenous fibers and other components of extracellular matrix (ECM). (Plopper and Hyde

2015).

Figure 2. 2 Illustration of various cellular shape and organization of epithelial

cells.


Muscle tissues of the airways mainly consists of smooth muscle tissues. These tissue types are

responsible for contractility of airways and are important in regulating airway size and resistance

(Plopper and Hyde 2015).

2.2.2 Tissue Morphology of the Trachea

The trachea has four layers of membranous tissues: mucosa, submucosa, cartilage and muscle,

and the adventitia. The mucosa, which forms the inner wall of the trachea, contains a ciliated

pseudostratified columnar epithelium layer with interspersed mucus-producing goblet cells that

warms, moistens and removes foreign particles that pass through the trachea (Paralta 2016).

Adjacent to the mucosa is the submucosa composed of loosely connective tissues, some blood

vessels, neurons (nerve cells) and glands. The glands in this tissue layer are called seromucous

glands because they secrete a mixture of water and mucus through narrow ducts to the luminal

surface where it is combined with the mucus produced by the goblet cells (Celis 2016). External

to the submucosa is a tissue layer consisting of cartilage that keep the airway open, and smooth

muscle fibers that contract during coughing which narrows the tracheal lumen. This narrowing

increases velocity which helps extricate the mucus containing foreign particles (Hlastala and

Berger 2001). The outermost layer is the adventitia that interconnects the cartilage rings with

loosely connective tissues (Paralta 2016).

2.2.3 Tissue Morphology of the Bronchi and Bronchioles

Similar but in different arrangement to the trachea, the bronchial wall is made up of tissue layers

composed of mucosa, lamina propria, smooth muscle and submucosa. The bronchus mucosa

layer has the same cellular structure and arrangement as the trachea. However, with successive

progression of small bronchial tubes into the bronchioles, the epithelium transitions into ciliated

simple columnar structure to cuboidal structure (Paralta 2016). Furthermore, the cartilage

content decreases and eventually disappears at the bronchiolar level, which leads to highly


folded mucosa layer due to the loss of support provided by the cartilage. The smooth muscle

and elastic fiber content increases in the lamina propria region at the bronchiolar level (Paralta

2016).

2.3 Pathophysiology

Asthma, chronic obstructive pulmonary diseases (COPDs), and respiratory infections such as

pneumonia and tuberculosis have been identified as some of the top conditions that contribute

to the global burden of chronic respiratory diseases (Ferkol and Schraufnagel 2014). With

Figure 2.3 Basic histology of airway tissues. A. Trachea cross-section showing

major layers. B. Lumen of a tertiary bronchus with various tissue layers. C. Lumen of a bronchiole. D. Alveoli cross-section. Images adapted from (Paxton et al. 2003)


increasing efforts to control and prevent respiratory infections through vaccination and

medication, the focus from a treatment perspective has shifted to non-communicable diseases

like asthma and COPD, as they are currently rising to be the leading causes of death (Bloom,

D.E. et al., 2011). Risk factors for asthma and COPDs include both environmental factors, from

exposure to personal or passive indoor and outdoor air pollution, and host factors such as

gender, genetic predisposition and more. With every breath, the lung airways are subjected to

airborne environmental pollutants, allergens, smoke and other irritants that the airways must

defend via a complex array of mechanical and biological defense mechanisms. In this

subsection, to demonstrate the complex pathological responses of these non-communicable

diseases we will address a few everyday respiratory defense mechanisms and

pathophysiological processes that lead to the development of asthma.

2.3.1 Airway Defense Mechanisms

The upper and lower respiratory tracts protect the lung against invading pathogens through

physical barrier at first entry, followed by various physical and biological defense mechanisms.

During inhalation, the tortuous anatomy of the upper airway prevents the penetration of particles

and organisms greater than 10 µm into the lower respiratory tract (Celis 2016). The particles are

trapped in the mucous layers of the airways. The bifurcating structures of the lower respiratory

tract further prevents particles above 3 - 5µm to penetrate the terminal or respiratory bronchioles

and alveolar space (Celis 2016).

The mucociliary transport system helps to remove particles attached to the mucous layer, which

surrounds the inner surfaces of airway lumens, from the bronchioles to the trachea via

movement induced by the beating cilia of epithelial cells. These cilia beat more than 1000 times

per minute, which moves the mucous layer up by approximately 1 cm per minute (Paralta 2016).

Various protein compounds such as mucoglycoproteins and proteoglycans along with

phospholipids secreted by the epithelial cells and submucosal glands alter the physical


properties of mucus, which facilitates the movement of mucus upwards. Other compounds

released by the epithelium and underlying tissues to the mucous layer include: Immunoglobin A

(IgA), which are antibodies that neutralize toxins or viruses and block bacterial entry to the

epithelium; lysozyme, which breaks the glycoside bonds of the large peptidoglycan molecules

that form bacterial membrane; and lactoferrin or peroxide, both of which inhibit bacterial

proliferation and growth (Plopper and Hyde 2015).

Epithelial cells play crucial roles in tissue barrier functions. Each cell is attached to its

neighboring cells via several junctions, namely tight junctions, gap junctions and desmosomes,

which create a barrier that is nearly impenetrable to invading species. Tight junctions, located on

the apical surface, form an impenetrable barrier between the luminal surface and the

intercellular space, while desmosomes, located on the lateral or basal surface, are structures

that form junctions between adjacent cells giving mechanical strength to the tissue layer (Celis

2016). Gap junctions, which are intercellular channels formed between adjacent cells, allow for

the sharing of ions and defence molecules between cells. Together these junctions create

effective mechanical barriers and maintain ionic gradients for movement of molecules and other

substances between tissue layers (Celis 2016).

CD4+ and CD8+ T lymphocytes are two distinct white blood cells that play crucial roles in

cell-mediated response to intruders like bacteria and viruses by producing cytokines to activate

the body's immune response (Plopper and Hyde 2015). These cells are present in the bronchial

epithelium in the absence of inflammation. Other white blood cells that are also a part of the

immune system components responsible for combatting infections and parasites include mast

cells and eosinophils. Mast cells reside in the tissues near blood vessels beneath the

epithelium. Upon activation by antibodies produced by B lymphocytes as a reaction to allergens

or pathogens, these cells release mediators that cause characteristic symptoms of respiratory

illnesses like allergy and attracts inflammatory cells(Celis 2016). An example of the


inflammatory cells that mast cells attract is eosinophils, which typically circulate in the blood and

migrate to sites of inflammation and infection in response to cytokines produced by mast cells

and other cell types.

The alveolar space is not protected by cilia and mucus, as it would obstruct and slow down the

movement of oxygen and carbon dioxide through the air-liquid interface (ALI). The main defense

system at the alveolar level involves numerous resident mononuclear mobile cells called

phagocytes. The type of phagocytes located in the alveoli are called alveolar macrophages

(AMs), and these cells have the ability to migrate to and neutralize invading pathogens by

binding to them via surface receptors, ingesting or killing them if any are living, and digesting

them (Celis 2016). These macrophages can also initiate lung inflammation by secreting cell

signaling proteins IL-1α, IL-1β or TNF-α, leading to the activation of neighboring cells and a

cascade of further immune reactions. If AMs fail to control the invading species, additional

phagocytes such as neutrophils are recruited to ingest or kill the pathogen. Neutrophils, when

activated by cell signalling proteins IL-1α, IL-6 or TNF-α produced by AMs and other cells,

perpetuate the immunological initiation process by secreting its own cytokines, destructive

enzymes and other cytotoxic compounds that lead to the recruitment of more neutrophils and

other cell types (Celis 2016).

The airway immune system is complex and a comprehensive summary of all defense

mechanisms are beyond the scope of this review. Highlighted above are a few of the key

defense mechanisms, typically involved in airway defence against irritants and harmful

substances, that will help us understand the specific pathophysiology of asthma.

2.3.2 Asthma Pathophysiology

Although initial responses, as outlined above, to toxins and other invading organisms are vital

for inducing the appropriate immune response, prolonged and unregulated release of cytotoxic


compounds as a result of continuous exposure to allergens and pathogens, cause chronic

inflammation and can lead to acute, sometimes permanent tissue damage.

Patients with asthma exhibit airway obstructions or hyperresponsiveness that are characterized

by constriction of airways through the tightening of surrounding smooth muscle, and basal

membrane thickening that leads to the symptoms of coughing, breathlessness, chest tightness

and more (Lambert et al. 1993). Asthma is induced by allergens such as plant or animal proteins

and the occurrences are often correlated with increased hygiene. High hygiene level minimizes

exposure to allergen which prevent the body from strengthening its immune system. The initial

inflammatory response process in asthma encompasses the infiltration and activation of CD4+ T

lymphocytes, mast cells, neutrophils and eosinophils. Chronic inflammation is thought to

promote significant structural changes to the airway walls that lead to airway remodeling. Airway

remodeling results from numerous changes including hyperplasia and hypertrophy, the

enlargement and shrinking, respectively, of airway smooth muscle cells, ECM abnormalities,

fibrosis of the tissue underlying the epithelium, remodelling of the microvasculature, and

epithelial metaplasia where the epithelial cells convert into a different cell type in response to the

changing microenvironment.

It is evident from the above-mentioned summary that no single cell type contributes alone to the

progression diseases like asthma but rather it is the cell-to-cell communication at various tissue

levels that highlights the complex nature of the inflammatory process. Thus, elucidating the

various diseases mechanisms and organ level responses affecting the airways require an

understanding of the collective impact of surrounding tissues and organ structures.

2. 4 Lung Tissue Models: In Vivo vs. In Vitro

Models in biology play essential roles as research tools in tissue engineering and clinical

applications. They facilitate the in-depth and precise study of various biological processes


including tissue repair, basic tissue functions and tissue responses to drugs and invading

species. However, as briefly alluded to before, accurate modeling of functional organs or tissues

requires examination of tissues, organs and organ systems using a more holistic and

whole-body approach. This is especially important for pathogenesis studies of respiratory

pathogens.

Due to the complex pathophysiology of respiratory diseases, inhalation toxicity and drug

response studies involving the assessment of airway reactivity and bronchial responsiveness

were and still continue to be predominantly conducted in animal models. Typically, these

models, which include mice, rats, guinea pigs and rabbits, are either developed to mimic

aspects of various respiratory disorders or they exhibit many of the same types of pathologies

as humans in response to disease-relevant agents (Chinoy, Antonio-Santiago, and Scarpelli

1994). One major caveat of this approach, however, is that some agents only function in certain

models and the appropriate stimulant, concentrations and drug standards found may be strain

or species dependent; as such, these models are often not predictive of human conditions.

Although the convergence of genomics and molecular pharmacology with bioinformatics has

helped improve our understanding of animal models and apply this understanding to human

conditions, the high cost and time considerations of whole animal studies are still prevalent

issues.

In light of this, in vitro models have been developed that reproduce many known components of

human respiratory responses. Compared to animal models, classic in vitro models not only offer

the ability to conduct high throughput cell culture studies that are relatively inexpensive, but also

provide information on specific cell behavior and interactions with relevant human cell types.

While classic cell culture models have been used to gain insight into cell and tissue-based

responses, physiologic functions and pathologic changes of the human respiratory system, the

influence of more complex structural elements and the physical environment found in vivo often


remains unaccounted for. The development of improved systems involving specialized 2D

chambers of 3D human lung constructs have allowed us to selectively add specific cell types in

order to study specific physiological interactions within the lung. Thus these systems simplify the

examination of the roles individual cells and their cellular mechanisms along with immune

factors involved in lung responses.

Recent advancements in microfluidic-based model development have improved upon current

three-dimensional, multicellular in vitro modelling techniques (ie ALI co-culture), by

incorporating relevant mechanical microenvironment and spatiotemporal chemical gradients of

the lung, discussed later in this section. Nevertheless, many challenges still exist in developing

models of complex tissues structures such as the ones found in the lung. Accurate modelling of

the respiratory tract, including structures from the trachea to the distal lungs, will need to reflect

the cell phenotypes, morphological structures and functional features of these regions. Thus to

study physiological responses of the lungs and disease development in these respiratory

regions, it is important to determine the necessary parameters that must be reconstituted.

A typical functioning airway tissue has: 1) a well-formed ciliated epithelium with proper barrier

functions depending on region along the tracheobronchial tree, 2) supportive scaffolding, ECM

structures and cell or tissue types, including smooth muscle, below the epithelium, 3) and the

underlying vasculature serving nutrients and oxygen delivery functions. Incorporation of these

factors into an in vitro model will include the selection of appropriate cell types with reproducible

generation and maintenance of specific biological functions, the development of relevant

scaffolds and matrices that support three-dimensional orientation and growth of tissues and

appropriate culture systems that are designed to accurately reflect the morphological structures

and the physicochemical properties of the airway microenvironment.


In this subsection we will highlight key factors to consider in the quest towards developing more

physiologically relevant in vitro models along with the strengths and limitations of existing

approaches taken to address these factors.

2.4.1 Selection of Cell Types and Cell sources

A major concern in developing more complex in vitro systems is the selection of appropriate cell

sources and cell types that can generate and maintain the tissue-specific biological functions of

an organ while in culture with other cell types. Mature cell lineages of the lung include ciliated

epithelial cells, Clara cells, smooth muscle cells, endothelial cells, and specialized pneumocytes

(Andrade et al. 2007). Lung primary alveolar epithelial cells (AECs) and human

tracheal-bronchial cells have been widely used in developing engineered lung models, for their

accurate demonstration of in vivo behavior (Mondrinos et al. 2006). These cell types are often

used as the gold standard to compare and validate models with other cell types. Problems, in

general, with the use of primary cell types are related to difficulties in obtaining and maintaining

viable human tissues and subsequently isolating a large number of viable cells. Alternatively,

transformed cell lines such as human submucosal adenocarcinoma cell line (Calu-3), human

bronchial epithelial cell line (16HBE14o) and human lung adenocarcinoma alveolar basal cell

line (A549) are often used to produce simple physiologic models of the lung because they are

able to express transport systems that are also present in human airway in vivo in a

reproducible manner (Andrade et al. 2007). However, disadvantages of using transformed cell

lines include the loss of normal cell responses due to the fact that these cells are isolated from

human cancers and therefore grow at an increased rate and divide many more times than a

normal human cell. As such, selection of cell types in model development will depend on the

type of model being constructed (pathological or physiological).

For instance, when modeling physiologic responses to lung irritants or viral agents, it is

important to consider the phenotypes of cells that contribute to the barrier functions of the airway


and permit entrance and replication of viral agents. Influenza virus, for example replicate in the

ciliated epithelium of the respiratory tract. Thus, to produce a 3D upper respiratory model to

study this effect, differentiated cultures of human bronchial tracheobronchial epithelial (HBTE)

cells should be used. HBTE cells are known to produce pseudostratified, ciliated and polarized

epithelium, which resemble the airway epithelium (Mondrinos et al. 2006). Furthermore, the

epithelium should be functionally similar to normal human tissues by producing differentiated

secretory and basal cells that contribute to disease susceptibility.

On the other hand, when developing pathological models for lung diseases like asthma, it is

important to consider factors that contribute to airway obstructions and airway

hyper-responsiveness, which are prevalent in these diseases. In the case of asthma for

instance, smooth muscle cells are known to be the main effector cells for airway narrowing,

caused by chronic inflammation in the lung that induce contraction of airway smooth muscle

(ASM) and changing the structural components of the airway wall including the ASM (Ebina et

al. 1993). Therefore, to model asthma, primary ASM cells or primary human bronchial smooth

muscle cells (HBSMCs) should be incorporated into the design for their contractile phenotype

that can be used to model contraction of ASM.

Aside from the section of cell types, a sustainable cell source is also imperative for

standardization of a lung model. In the past, human embryonic stem cells (hESCs) and induced

pluripotent stem cells (iPSCs) have been shown to generate lung epithelial cells. Additionally

murine ESCs have been used to produce conducting airway tissues including ciliated epithelial

cells, Clara cells and surfactant producing type II AECs (Nichols, Niles, and Cortiella 2009).

Although, not many reports have demonstrated the use of these differentiated cells or lung

tissues in the development of physiologic lung models, the generation of cells from renewable

sources, such as hESCs, hESCs and murine ESCs, is appealing for the production of replicate

cultures and standardization of lung models.


2.4.2 Selection of Physiologically Relevant Scaffold

The primary requirements for any scaffold used in regenerative medicine practices are

biocompatibility and the ability of the material to provide for 3D development of tissues. For lung

tissue development, the scaffolding must retain its structural integrity long enough to provide a

necessary framework to support cell growth and tissue development without impeding

characteristic tissue properties, such as the organization of cells in tissues. The biomaterial

must also be as elastic as normal tissues so as to not induce any restrictive conditions that may

resemble an abnormality such as the thickening of connective tissues in fibrosis. Porosity is also

important for the transfer of nutrients and waste removal from tissues. The scaffold must also

have adequate chemical compositions to promote cell attachment and interactions or cell-cell

signalling with other cells types in 3D space as found in vivo. To meet all of these requirements,

a hybrid scaffold formed from various materials may need to be designed.

In the past, both natural and synthetic polymers have been used in engineering lung model

systems with various degrees of success. Many natural hydrogels have been produced that are

chemically and structurally similar to the ECM found in the body. Since biophysical and

biochemical cues generated by the in vivo ECM along with their mechanical properties provide

major signaling sources for regulating fundamental cell behavior as well as growth and

differentiation of cells (Samadikuchaksaraei et al. 2006), these hydrogels are commonly used in

tissue engineering and model development. Types of natural hydrogels include collagen (Choe,

Tomei, and Swartz 2006), (Miller, George, and Niklason 2010), Matrigel (Mondrinos et al. 2006),

(Chinoy, Antonio-Santiago, and Scarpelli 1994), (Hirst, Twort, and Lee 2000), Gelfoam

(Andrade et al. 2007) , Pluronic F-127 (Cortiella et al. 2006) and Englebreth-Holms tumor

basement membrane (Blau et al. 1988). Some limitations in using these natural scaffolds

include weakened mechanical properties and variation in degradation rates (Nichols, Niles, and

Cortiella 2009).


Synthetic polymers on the other hand tend to have a much wider range of mechanical properties

than that of natural materials. To maintain the elastic nature of the environment, synthetic

matrices such as polyglycolic acid (PGA) (Badylak 2002), poly(lactic-co-glycolic acid) (PLA)

(Badylak 2002) and polydimethylsiloxane (PDMS) (Huh et al. 2010) have been used in lung

model development in the past. Although synthetic matrices can provide the mechanical

properties required to generate the elastic nature of some lung tissues, it lacks the chemical

complexity of ECM found surrounding these tissues.

Ideally the development of physiologic models of the lung involves the use of acellular (AC)

natural scaffolds, which are obtained from whole animals and are composed of ECM secreted

by the resident cells of the lung. This has been accomplished before in numerous studies, in

which AC whole mouse-trachea or AC human-lung are repopulated with mouse or rat fetal lung

cells, murine embryonic stem cells (mESCs) and rat type II AECs to examine the influence of

cell attachment and morphology. In one of the studies, where an AC rat lung matrix was

repopulated with mESCs, mESCs differentiated into ciliated epithelial cells, Clara cells, type II

AECs, endothelial cells and smooth muscle cells. Other studies have demonstrated the

reproduction of some of the morphological characteristics of alveolar epithelium. Although AC

scaffolds may provide significant contributions to the field of organ transplantation, the high cost

and accessibility of these scaffolds may impede its progress in model development.

2.4.3 Microfluidic Lung Models (Organ-on-a-chip systems)

In model development, organ-on-a-chip systems, as facilitated by advanced microengineered

technologies and microfluidic principles, have enabled the in vitro studies of human physiology,

disease progressions and complex tumour microenvironment in an organ-specific context.

Briefly, this is done by leveraging advanced microfabrication technologies to recreate key

aspects of the in vivo microenvironment, such as tissue-tissue interfaces, spatiotemporal

chemical gradients and the mechanical microenvironment that influence tissue behavior (Huh,


Hamilton, and Ingber 2011). More importantly, by enabling compartmentalization, this emerging

platform helps us improve our understanding of cellular and tissue behavior arising from their

interactions with multiple cell and tissue types (Moraes et al. 2012). In studies involving the

complex tumor microenvironment, these techniques have been used to model tumor spheroids,

tumor-stromal interactions, angiogenesis, oxygen gradients and intravasation (Young 2013).

Examples of these systems have previously been reviewed and will not be discussed at length

here with the exception of a few brief remarks on microfluidic lung models.

Current microfluidic lung models described in the literature focus on the impact of engineering

principles such as shear stress, mechanical strain, and surfactant-induced airway tensions on

lung tissue development. Many of the microfluidic platforms developed so far for modelling the

respiratory tract mimic the alveolar–capillary interface, and more specifically the epithelial and

endothelial barrier. We highlight a few examples of these lung models here to illustrate impact of

microfluidics on the development of a more physiologically relevant model. A similar design

feature present in all systems described here is the incorporation of a thin porous membrane

coated with collagen gel placed between two vertically stacked microfluidic chambers. Cells are

cultured as monolayer on these collagen-coated membranes and are believed to provide an

ECM-like interface between tissue layers.

An early platform developed by Huh et al. modelled cellular level lung injury caused by liquid

plug flows in a microfluidic system (Huh et al. 2007). The design involved a compartmentalized

air-liquid two-phase microfluidic system which consisted of two stacked PDMS chambers

separated by a thin polyester membrane (Figure 2.4 A-C). Small airway epithelial cells (SAECs)

were grown as monolayers on the polyester membrane with perfusion culture (media in both top

and bottom PDMS chambers) at first, followed by culture at ALI where the media was removed

from the top chamber to better resemble in vivo physiological conditions. The microfluidic

system was attached to a liquid plug generator that created and passed liquid plugs over the


epithelium monolayer and ruptured at a downstream location. This resembled the

surfactant-deficient reopening of closed airways in vivo that lead to epithelium injury. The study

demonstrated, the ability of the system to sustain long-term cultures of SAECs beyond 6 days

and for the reconstructed epithelium to experience injuries as a results of liquid plugs rupturing

on top of the epithelium monolayer. Furthermore, these result showed how microfluidic systems

can be used to induce fluid mechanical stresses similar to the conditions experienced by cells

and tissues in vivo.

Using a similar setup, an alveolar-capillary interface of the human lung was reconstructed

cytokines by Huh and colleagues that reproduced more advanced organ-level responses to

bacteria and inflammatory (Huh et al. 2010). The system consisted of two vertically stacked

PDMS chambers separated by a thin porous flexible PDMS membrane. Human AECs and

human pulmonary microvascular endothelial cells (HPMECs) were cultured and grown to

confluence on opposing sides of the PDMS membrane, coated with fibronectin and collagen.

The device also consisted of two side chambers that were attached to a vacuum to induce cyclic

mechanical stretching of the PDMS membrane to mimic physiological breathing movements

(Figure 2.4 D). This provided new mechanistic insight into the role that breathing plays on

cellular uptake of airborne particulates simulated by silica nanoparticles. The combination of

fluid flow in the vascular channel, the ALI created by exposing the epithelium to air and

application of cyclic mechanical strain strongly promoted differentiation of the epithelial and

endothelial cells. This was confirmed by enhanced surfactant production and vascular barrier

function, measured by both transepithelial electrical resistance (TEER) and assays of

macromolecular transport. The model demonstrated similar effects of breathing on the

nanoparticle adsorption as observed in a mouse ex vivo ventilation perfusion model, in their

system where the epithelial and endothelial uptake of nanoparticles and transport to the

underlying vasculature increased when the system was subject to cyclic mechanical strain. In

addition, when immune activators TNF-α and E-coli bacteria were introduced to the epithelium


layer to mimic inflammation, the endothelium was activated, as measured by increased

expression of surface intercellular adhesion molecule 1 (ICAM-1), and recruited more human

neutrophils when introduced to the vascular channel under flow conditions. The neutrophils

migrated through both the endothelium and the epithelium where they engulf E coli. An inactive

endothelium was observed when the neutrophils were introduced with the presence of TNF-α

and E-coli.

This lung-on-a-chip device was also used to model the dose-limiting side effects of cancer drug

interleukin-2 (IL-2), particularly, the induction of pulmonary vascular permeability leading to fluid

accumulation in the lung (pulmonary edema) and fibrin clot formation (Huh et al. 2012). IL-2 at a

clinically relevant dosage along with prothrombin and fluorescently labeled fibrinogen, which are

human blood plasma proteins, were perfused through the vascular channel over a period of 4

days. Formation of blood clots and the compromise of oxygen transport occurred over the same

2 to 4-day time course as observed when IL-2 was administered in patients. The study also

revealed that mechanical forces of breathing motions contribute to the development of

increased vascular leakage and subsequent pulmonary edema induced by IL-2. The results

from both of these studies highlight the value of microfluidic systems and their ability to

independently incorporate and vary cell types as well as induce both chemical and mechanical

stimuli to its tissue cultures.

A more recent example of a microfluidic-based lung model that uses the same ALI design,

(Figure 2.4 E), and demonstrate the influence of tissue – tissue cross talk is the human

small-airway-on-a-chip. In this study, primary human bronchiolar epithelium and pulmonary

microvascular endothelium was reconstituted in this device and exposed to a viral mimic,

polycytidylic acid (poly(I:C)) (Benam et al. 2016). Proinflammatory responses, including a lager

increase in inflammatory cytokine (RANTES, IL-6, and IP-10) secretions were observed similar

to that observed in humans with acute or severe asthma. The secretions were higher when the


bronchiolar epithelium was co-cultured with the vascular endothelium than when stimulated by

poly(I:C) in monoculture. However, the endothelium did not affect the secretions of other

inflammatory mediators (e.g., GRO-α, IL-8). E-selectin and vascular adhesion molecule 1

(VCAM-1) expression in the endothelium stimulated by poly(I:C) increased adhesion, rolling

along endothelium surface and recruitment of circulating human neutrophils introduced into the

vascular channel via perfusion.

In the same study, the device was used to model a few aspects of asthma and COPD. Cytokine

IL-13 exposure to the bronchiolar epithelium showed similar tissue behavior (goblet cell

hyperplasia, increased cytokine secretions and decreased ciliary function) to that seen in

asthma patients. The device was also used to culture COPD patient derived airway cells where

a differentiated COPD epithelium was formed and exhibited COPD phenotype. Additionally, the

COPD epithelium exhibited enhanced inflammatory responses induced by poly(I:C) compared

to the normal epithelium. Thus, the results of this study revealed the importance of crosstalk

between human lung airway epithelium and endothelium during inflammation, as facilitated by

incorporation of ALI for co-culture and the application of vascular perfusion within the

microfluidic chip.

Although the design of the microfluidic models discussed so far allowed it to be used in various

applications and provide insights about cellular responses and cell–cell interactions when

subjected to different stresses, there are limitations to the use of this platform.

One of these limitations is that results obtained from these chips may not necessarily be

comparable to human physiological responses even though primary human cells are used. This

is because cells are cultured on thin ECM-coated porous synthetic membranes, which may

provide the mechanical support for co-culture but lack the chemical composition and structural

organization of the native ECM. Cellular behavior and responses impacted by this change may

be misleading when compared to their responses in vivo where the structural microenvironment


influences reactions. Additionally, although the co-culture experiments in the small airway

reconstruction have provided valuable insights about pathologic responses of the airway in the

case of asthma and COPD, the tissue compositions of the small airways in the lung are very

different from a simple epithelium and endothelium and therefore, may impact tissue responses.

We know from previous reports that it is not only the epithelium but also the ASM tissue that

contribute to the expression and secretions of pro-inflammatory cytokines and mediators in the

pathogenesis of asthma and COPD. Thus to develop a more representative model of small

airway and its inflammatory responses in asthma and COPD the influence of smooth muscle

tissue must also be considered.


Figure 2.4 Microfluidic in vitro airway models. (A-C) The microfabricated small

airways are comprised of PDMS upper and lower chambers sandwiching a porous membrane. SAECs are grown on the membrane with perfusion of culture media in both upper and lower chambers until they become confluent. Once confluence is achieved, media are removed from the upper chamber, forming an air–liquid interface over the cells. Source: (Huh et al. 2007).D) Lung-on-a-chip microfluidic device with compartmentalized PDMS microchannels to form an alveolar-capillary barrier on a thin, porous, flexible PDMS membrane coated with ECM. The device recreates physiological breathing movements by applying vacuum to the side chambers and causing mechanical stretching of the PDMS membrane forming the alveolar-capillary barrier. Source: (Huh et al. 2010). E) Small-airway-on-a-chip design. Source: (Benam et al. 2016).

D

E


2.5 Thesis Objectives

Organ-on-a-chip systems incorporate 3-D microstructures, basic tissue components and

simulate the mechanical and physiological responses of entire organs in their design. Progress

to date has been substantial, with the reproduction of complex mechanical microenvironment of

healthy and diseased tissues of several major organs leading to similar tissue responses to the

in vivo state. Several reports have claimed partial functionality of specific tissue types and

demonstrated communication between different tissues reproduced in these systems. Recently

there is a strong interest in combining multiple organs on a single chip to demonstrate the

potential for a human-on-a-chip platform. Although the approaches in designing organ-on-a-chip

systems vary and is dependent on the research question, the common goal among researchers

is to create systems that resemble the complex in vivo microenvironment in a more

physiologically relevant way than current technologies permit.

The main goal of this thesis is to develop and perform preliminary assessment on a

microfluidic-based in vitro model that mimics the in vivo tissue microenvironments of the human

airway, in a more physiologically relevant way with respect to cell-cell and cell-ECM interactions.

We suspend inside a thermoplastic microfluidic device a layer of ECM like gel that resembles

the chemical composition and structural complexity of the extracellular space in airway tissues,

and incorporate a co-culture of two airway cell types, that play vital roles in inflammatory

responses of the airway (Figure 2.5). An epithelial cell line called Calu-3 and primary human

bronchial smooth muscle cells (HBSMCs) are used in this model. Preliminary assessment of the

in vitro model is conducted to demonstrate basic tissue characteristic, found in the native airway

epithelium and smooth muscle tissue. In addition, the aim of this thesis is to integrate the system

into an arrayable platform for more efficient culture experiments and higher-throughput

co-culture studies of airway tissues. Successful development of this model would provide an in

vitro system for conducting long-term studies on the physiology and pathology of the lung

airway.


To achieve this overall objective, the specific aims of this thesis are as follows:

1. To devise a protocol for fabricating and assembling an array of airway-on-a-chip systems that

can support co-cultures of airway epithelial and smooth muscle cells. Each device should have

an array of compartmentalized units that are isolated from other systems. Each system should

include separate compartments for the culture of the two airway cell types.

Figure 2.5 Simplified model of the airway wall tissue structure involving the airway epithelium surrounded by a thin layer of ECM and a band of aligned smooth muscle tissue. The device design will consist of two vertically stacked compartments with a top air flow chamber and bottom media reservoir to support culture of airway Calu-3 epithelial cells and HBSMCs, respectively.

2. To develop an analytical model that can predict device design parameters for facilitating

suspended ECM hydrogel flow inside the device. The analytical model should be validated by

experimental data.

3. Investigate cell adhesion behavior on ECM hydrogels and determine cell culture parameters

for long-term culture (> 3 weeks) inside the device. An investigation of cell adhesion to ECM like

gels, such as collagen type 1 and Matrigel or a mixture of these gels, should be conducted. It

smooth muscle tissue

epithelium Lumen

extracellular matrix (ECM)


should include an assessment of initial cell seeding densities of each cell types, inside the

device, for a 2 to 3-week cell culture period.

4. To perform preliminary assessment of basic tissue characteristics in order to assess the

fitness of the device as an airway model. Staining and imaging protocols should be well

developed to effectively analyze and evaluate tissue characteristics and conduct studies on

culture conditions that affect tissue formation. Calu-3 epithelium tissue barrier functionality and

HBSMC alignment in both monoculture and co-culture systems should be evaluated.

2.6 Future Outlook

Classic in vitro platforms (eg. multi-well culture plates) over the past few decades have served

as an important laboratory research tool in the area of in vitro model development. However, the

progress made by organ-on-a-chip technologies over the past decade, especially in the context

of a complex organ system like the lung has revolutionized the way we approach in vitro model

development. The goal of in vitro model development, in the near future, is to create systems

with specific genetic profiles of individuals to assess the significance of single gene products

and pathways in respiratory illnesses. To do this, current organ-on-a-chip technologies will not

only need to support high throughput cell culture studies but also be easily accessible and

adaptable to its end users.

29

3.0 Airway-on-a-Chip Design and Fabrication

With growing interest and increased efforts of using microfluidics as a research platform, more

tools and fabrication methods are being used for developing high volumes of microfluidic

systems in a more efficient manner. Plastics have an unmatched popularity when it comes to

compatibility with biological applications (due to their favorable properties such as inertness and

low molecular adsorption (Berthier, Young, and Beebe 2012), and compatibility with existing

manufacturing processes that make them a low-cost material suitable for mass production. For

these reasons, researchers are dedicating efforts to translate microfluidic research commonly

done in PDMS, to plastics, especially for those interested in commercializing their designs.

Many plastic fabrication methods are available for researchers and depending on the stage of

technology development and the technical needs, these fabrication methods are more or less

suitable. The advantages and limitations of each fabrication method are reviewed elsewhere,

(Becker and Locascio 2002) and therefore will not be discussed at length here. For early stages

of technology development, where a short design-to-prototype time is desired, micromilling is a

well-developed, low start-up cost, low-volume producing fabrication method that meets the

needs of researchers (Guckenberger et al. 2015). It is a fabrication method that removes bulk

material (usually metal or plastic) from a starting stock piece, using a rotating cutting tool, to

create microscale features. The milling of features with precision and in a repeatable fashion is

controlled by an automated computer numerical control (CNC) system, which translates

computer-aided design (CAD) to its finished parts. This method has proven to be very effective

3.0 Airway-on-a-Chip Design and Fabrication 30

in many recent developments of biology-based plastic microfluidic systems. For instance, plastic

micromilled devices have been used to conduct a variety of assays in the Kit-On-A-Lid- Assay

system (E. Berthier et al. 2013), for cell capture and isolation of DNA, RNA and proteins

(Strotman et al. 2013) and even to study the microenvironment of a co-culture system involving

primary fetal testis cells (Carney et al. 2014). Given the success of these micromilled plastic

devices along with ultra-rapid prototyping capabilities of this technique compared to soft

lithography (Guckenberger et al. 2015), we have chosen micromilling as our fabrication method

to build our airway microfluidic in vitro model.

For biological applications, plastics such as polystyrene (PS), poly methyl methacrylate (PMMA)

and cyclin olefin copolymer (COC) are commonly used as cultureware and are more reliable to

biologists on account of the decades of validated research provided by these materials. In

addition, PDMS, in the context of cell culture and assay studies, have been shown in some

cases to induce adverse effects on culture systems due to leaching of uncrosslinked oligomers,

that may interact detrimentally with the culture, and adsorption of hydrophobic molecule

introduced to the system, which may impact assay results. In comparison thermoplastics do not

leach under culture conditions and has minimal affect on absorption of small molecules.

Because our device will have compartmentalized units, bonding of multiple thermoplastic layers

will need to be considered in the microfabrication process of our device. Bonding of PMMA is

well documented in the literature, with respect to microfabrication, compared to PS, because

polystyrene deforms well below its glass transition temperature making it difficult to retain

microscale features under thermal-assisted bonding conditions. This improves upon the

manufacturability of PMMA and is likely the reason why it is preferred over PS in general

plastic-based applications. We have chosen to fabricate our devices using off-the-shelf PMMA

for convenience. we expect to see similar biological results obtained from devices made out of

PS as we will see from PMMA.


3.1 Microfluidic Design

The microfluidic design of our device will include two vertically stacked compartments for each

of the airway cell types, separated by an ECM hydrogel suspension, which will accommodate

the co-culture of both cell types (Figure 3.1A, B). There will be one separate inlet and one

separate outlets for each of the cell types to accommodate media changes from each cell type

to replenish nutrients during long term culture (Figure 3.1C). Since inlet and outlet ports for

Calu-3 cells are located above the hydrogel, to avoid disturbing the hydrogel during media

chances and staining protocols, the ports should be located away from the region where the

ECM hydrogel will be suspended (Figure 3.1 D).

3.2 Device Microfabrication and Assembly

As previously mentioned, to fabricate our devices we will be using micromilling to carve out

microscale features into the layers of our device (Figure 3.2 A). After milling, the layers of

PMMA will be bonded to build compartmentalized units to culture two different airway cell types.

The bonding technique used will be a solvent-assisted thermal bonding for PMMA, well

documented in the literature. A solvent such as ethanol and isopropyl alcohol is introduced

between two layers of PMMA and pressed together using a Carver hydraulic press at a

temperature below the glass transition temperature of PMMA (Figure 3.2 B). Because of the

decreased solubility parameter of PMMA at elevated temperature, the solvent is able to assist in

bonding two PMMA pieces together below its glass transition temperature. The solvent used for

our case will be ethanol, and operating conditions will be at 70oC with 1000 lbf.


Figure 3.1 PMMA microfluidic device for co-culturing airway cells. A) Three

layers of PMMA will be used to build a device with two vertically stacked microfluidic compartments separated by a thin hydrogel layer. B) The hydrogel will be suspended inside the microfluidic device to culture Calu-3 epithelial cells on top of the hydrogel and HBSMCs on the underside. Culture media will be introduced to both the top and bottom compartments during culture. To create an ALI the media from the top compartment will be removed once a stable Calu-3 epithelium has formed on top of the hydrogel. C) Two separate inlets and outlets are designed into the system for seeding and maintenance of each cell types in culture. D) The outlet port for the top compartment (Calu-3 cells) is located away from the suspended hydrogel to avoid disturbing the hydrogel during culture.

Inlet port: HBSMC cell seeding

Inlet port: Calu-3 cell seeding

A

B

C D Outlet

ports

Outlet port:


3.3 Materials and Methods

3.3.1 Device Design and Fabrication

Devices were designed and modeled in Solidworks 3D modeling software (Dessault Systems,

Velizy-Villacoublay,France) and fabricated using a plastic micromachining method.

Microfeatures modelled in Solidworks were imported into SprutCAM (SprutCAM, Naberezhnye

Chelny, Russia) to be converted into TAP files for milling on CNC milling machine. Device layers

were fabricated with a Tormach PCNC 770 vertical milling machine (Tormach, Waunakee, WI),

from clear 1.5 mm or 2.0 mm thick sheets of poly (methyl methacrylate) (PMMA, McMaster Carr

Supply Company, Elmhurst, IL, USA). Devices were milled using 1/16” (1.5875 mm) (Part #

01982156), 3/64″ (1.1906 mm) (Part # 07765431) and 0.015″ (381 μm) (Part # 37289501)

endmills.(MSC Industrial Supply Co., Melville, NY, USA).

Carver Hydraulic Press

Bonding at 70

0 C &

1000lbf

CNC Milling Machine

Micromilling

Solvent Assisted Thermal

Bonding

Plate

Plate

Steel

Plate

s

A B

Figure 3.2 Microfabrication of PMMA microfluidic device for co-culturing airway cells. A) Microscale features are milled into a sheet of PMMA using a CNC

milling machine to fabricate the three layers of the device. After milling PMMA chips are cleaned and prepared for bonding. B) PMMA chips are aligned with a thin film of ethanol (99%) between each chips and bonded using a Carver hydraulic press.

PMMA Layer

PMMA Layer

PMMA Layer

PMMA Layer

PMMA Layer

PMMA Layer

PMMA Layer

PMMA Layer


The top layer

A 2.0 mm thick PMMA sheet was used to fabricate the top layer. It consisted of a series of

rectangular channels with curved edges and through holes. Each channel had two through

holes, an inlet and outlet for one cell type, located inside at each short edge of the channel, and

two through holes located 1.25 mm outside each short edges of the channel (Figure 3.3 A). The

channels were 1.2 mm tall, 5 mm in diameter and spanned 10 mm from the centre of the two

through holes located inside edges of the channels. The inlet and outlet through holes were 2.00

mm and 2.75 mm in diameter respectively.

The middle layer

The middle layer was fabricated using a 1.5 mm thick PMMA sheet and consisted of channels

and the outside through-holes in the same orientation and dimensions as the top layer. The

channel depths were adjusted to 0.7 mm. A second set of smaller rectangular channel, with

curved edges were inserted inside each larger rectangular channel. The smaller channels were

0.65 mm deep, 2.25 mm wide and 6 mm in length. The inlet edges of the smaller channels were

mirrored with the inlet through-holes located inside the edges of the top layer. A set of smaller

rectangular through-holes with curved edges were designed inside the smaller rectangular

channels with dimensions 0.65 mm X 2.0 mm (Figure 3.3 C).

The bottom layer

The bottom layer of the chip consisted of an array of inlet and outlet ports connected by an array

of straight-channels that are 12.5 mm long and 1.6 mm wide. Both the channels and the ports

were 1.00 mm deep. The inlet and outlet ports were 2.75 mm and 2.0 mm in diameter

respectively and were aligned with outside inlets and outlets of the top layer. The straight

channels were aligned at the center with the larger channels of the middle layer (Figure 3.3 B).


After milling, each layers of the device was cleaned with laboratory soap and water, rinsed in

70% ethanol in DI water to remove any residual PMMA flakes and coolant from the milling

process. The layers were subsequently dried with dry compressed air and any remaining PMMA

pieces were removed using FisherbrandTM Label Tape (Fisher Scientific, Waltham,

Massachusetts, USA). See Appendix A5 for a more detailed protocol.

3.3.2 Solvent-Assisted Thermal Bonding

The cleaned PMMA layers were bonded using a thermal-assisted thermal bonding method. 99%

ethanol (Commercial Alcohols Inc., Toronto, ON, Canada) was applied via pipetting in between

two layers of PMMA and the layers were pressed together to create a thin film of solution

between the layers. First, the top and middle layers were aligned to mirror each other and the

PMMA stack was placed between two blank steel plates, which were pre-heated to 70oC

(Figure 3.2 B). A force of 1000 lbf was applied to the steel layers and the stacked PMMA with a

bonding temperature of 70oC for 1 min in a Carver Automatic Hydraulic Laboratory Press

(Carver Inc., Wabash, Indiana, USA). The bonded PMMA stack was removed from the hydraulic

A B

Figure 3.3 Schematic of device design and parameters for a 6 system PMMA co-culture device. Top view of the top A), bottom B) and C) middle PMMA layer. D)

Cross-section of one bonded system.

C D


press and cooled down to room temperature. The bonding solution was then pipetted in

between the stack and the bottom PMMA layer, which was aligned so that the milled features of

the bottom layer mirrored the un-bonded face of the middle layer. The three-layer stack was

then placed between two blank steel plates and the bonding process was repeated as described

above. After bonding and subsequent cooling of the PMMA stack, the remaining liquid was dried

with dry nitrogen gas and left at room temperature overnight to all existing volatiles to evaporate.

3.4. Results

After microfabrication and bonding, devices where ready for sterilized and loaded with a

suspended ECM hydrogel. The devices can be designed to have any number of systems

desired for a particular test. Figure 3.4 shows two examples of assembled devices with 6

systems in one and 8 systems in the other.

Figure 3.4 Photograph of assembled PMMA microfluidic device. Channels highlighted in blue represent the suspension channels for ECM hydrogel (A, C). Channels highlighted in green (B) represent the space where Calu-3 epithelial cells will be cultured HBSMCs are cultured in the channels highlighted in red (D).

A

B C D

37

4.0 Suspended Microfluidic Principles

One of the key design aspects of our microfluidic design is the suspended ECM hydrogel that

will accommodate the co-culture of Calu-3 epithelial cells and HBSMCs on its opposing

surfaces. To closely mimic the dimensions of the ECM tissue-tissue interface typically found in

airway tissues, we will need to design a system that can suspend an ECM hydrogel that is very

thin but has a large enough surface area that upon Calu-3 cell culture can be representative of

an epithelium monolayer (Figure 4.1). We should also aim to maximum the area of co-culture

region to build a more representative model of airway walls. Our goal for this chapter is to

develop an analytical model to help us predict device design parameters that will accommodate

the suspension of a thin ECM hydrogel, with large ALIs (exposed surface areas) for potential ALI

culture, and to maximize co-culture surface area.

To achieve this goal, we looked at capillary based microsystems that have the advantage of

moving fluid autonomously by way of a fluid dynamic phenomenon known as spontaneous

capillary flow (SCF). This is because, as per our current microfabrication protocol, the ECM

hydrogel loading into the device takes place after the device has been bonded and sterilized,

leaving us with no direct access to the suspension channel except at the inlet of the top

compartment. Therefore, we needed a technique that allows us to load ECM gel into the system

without having direct access to the suspension channel.

4.0 Suspended Microfluidic Principles 38

SCF is where surface tension drives the movement of fluid spontaneously along the surface of a

solid or absorbent material. Recently a self-propelling fluid handling technique known as

"suspended microfluidics” has been introduced for microfluidic applications requiring microscale

flow within open microstructures (Casavant et al. 2013). In these systems, SCF occurs within

microchannels with multiple air or immiscible fluidic interfaces. These interfaces are open to

allow direct access to the flowing fluid (in other words, having larger ALIs). An analytical

relationship describing the design conditions for SCF and their limits is as follows:

𝒑𝒇

𝒑𝒘< 𝒄𝒐𝒔 𝜽𝒔𝒐𝒍𝒊𝒅 𝒎𝒂𝒕𝒆𝒓𝒊𝒂𝒍 (4.1)

where pf is the cross-sectional perimeter of the free interface. pw is cross-sectional perimeter of

the wetted surface or solid material. In our case, pw is 2H. The θsolid material, is the contact angle of

a liquid against a surface, (ie., it is a measure of the wettability of a surface). The free interface is

surface that is exposed, which for our case presented in Figure 4.1 is equivalent to w, however

since there are two surfaces exposed, the pf is equal to 2W. It should be noted that the

Figure 4.1 Schematic of a simplified bronchiole wall (left) with a ciliated monolayer epithelium surrounded by a thin layer of ECM wrapped around by a band of smooth muscle tissue. Cross-section of our proposed model design (right) where

our aim is to maximize the co-culture region, represented by W, while minimizing the thickness of the suspended ECM hydrogel, represented by H.


theoretical limits of cosθ are -1 ≤ cosθ ≤ 1 which means that if the surface is easily wetted the

maximum value of cosθ would be 1 and if the surface cannot be easily wetted the value of cosθ

would be -1.

𝒑𝒇 − 𝒑𝒘𝐜𝐨𝐬 𝜽𝒔𝒐𝒍𝒊𝒅 𝒎𝒂𝒕𝒆𝒓𝒊𝒂𝒍 < 0 (4.2)

If we rearrange the above relationship as follows and apply the maximum cosθ value of 1 (ideal

situation where the liquid wets the surface very well), it states that in order for SCF to occur, the

difference between the free perimeter and the wetted perimeter must always be less than 0. In

other words, the free perimeter cannot be greater than the wetted perimeter and in our case it

means the thickness of the suspended hydrogel must always be greater than the width of the

hydrogel. This is the exact opposite of what we aim for our system to accommodate.

We noticed in our preliminary investigation, however, that we can flow hydrogel solutions into

our system where the height of the channel (thickness of the hydrogel) was much smaller than

the channel width (hydrogel width) as long as we loaded the hydrogel into the channel with a

pipette at a low speed. In the context of fluidic principles, this loading of hydrogel translates to a

pressure applied by a constant flowrate of fluid at the inlet of the open channel to help move the

fluid-front forward. We call this type of flow “pressure-driven flow” since the flow is driven by the

pressure applied at the inlet of the channel. This is in contrast to a self-propelled flow where no

pressure is applied, which is described as SCF. This can also be thought of from a force balance

perspective where for the SCF case, the capillary force is high enough to move the flow of fluid

forward and for the pressure-driven flow case an external pressure needs to be applied for flow

to occur. See details in Appendix A2.

In this section we describe the modifications made to the analytical relationship described

above, in equation 4.1 to accommodate the observations we have made in our system with


pressure-driven suspended microflow. A general analytical model that predicts the onset of

pressure-driven suspended microflow is presented here. See derivation in Appendix A1.

[𝒑𝒇

𝒑𝒘− 𝒄𝒐𝒔 𝜽𝒔𝒐𝒍𝒊𝒅 𝒎𝒂𝒕𝒆𝒓𝒊𝒂𝒍] 𝒑𝒘 𝜸𝑳𝑮

𝒅𝒙

𝒅𝑽< 𝑷 (4.3)

where γLG, represents the interfacial tension of the liquid, P represents the pressure applied at

the inlet of the channel, and dx/dV represents the inverse of the cross-sectional area of the fluid

front.

This analytical condition states that given a set of system parameters (left side of the equation),

flow will occur only when the applied pressure at the inlet is higher than the values obtained on

the left side of the equation. Since applied pressure is a user-defined parameter this gives the

user more flexibility where depending on all other system parameters the user can determine

the minimum pressure or flow rate required to induce flow happen. See Appendix A1 for

minimum pressure calculation.

Next we wanted to validate our proposed model with experimental data. To do this we chose a

simple system where pressure-driven flow occurs between two vertically parallel channel

walls,without a ceiling or a floor, where the heights of the channel walls are smaller than the

distance between them (ie. width/height aspect-ratio greater than 1). This case does not meet

SCF conditions which is why pressure or a constant flow rate will need to be applied at the inlet

of the channel (Figure 4.2).

For this system, based on force balance analysis of pressure-driven suspended microflow

between parallel walls, we derived analytical equations for (i) liquid penetration distance, x, with

respect to time, equation 4.4, and (ii) liquid penetration velocity, V, with respect to time, equation

4.5. See Appendix A2 for derivation.


𝑥(𝑡) = √[∆𝑝𝑤2

6𝜇+

𝛾𝑤

3𝜇[𝑐𝑜𝑠𝜃 −

𝑤

ℎ]] 𝑡 (4.4)

𝑉 = √∆𝑝𝑤 2

24𝜇+

𝛾𝑤

12𝜇[𝑐𝑜𝑠𝜃 −

𝑤

ℎ] √

1

𝑡 (4.5)

Δp represents the pressure difference between the inlet of the channel and the fluid front; w, h

are the width and height of the channel, respectively; γ and µ are the surface tension and

dynamic viscosity of the fluid; θ is the contact angle of the liquid with the solid surface; and t is

time. An experiment was conducted to assess whether the proposed models predict

experimental outcomes.

Figure 4.2 Schematic of pressure driven suspended microflow between parallel walls for channel width/height aspect-ratio greater than 1.

Pipette with fluid

Fluid flow thorough a channel

with no floor or ceiling.



4.1.1 Device Design

A 2.0 mm thick PMMA sheet (PMMA, McMaster Carr Supply Company, Elmhurst, IL, USA) was

used to fabricated a PMMA device with a 20 cm long suspended channel. The device consisted

of a large rectangular channel with (w = 2cm, h = 1.3mm, L = 22 cm) dimensions. Another

channel with (w = 900µm, h = 650µm, L = 20 cm) dimensions was embedded inside the larger

rectangle. The smaller channel had two ports, an inlet and outlet, located at each short edge of

the channel. The inlet and outlet ports were 2.00 mm and 2.75 mm in diameter respectively and

450 µm tall.

4.1.2 Micromilling

PMMA devices was designed and modeled in Solidworks (Dessault Systems,

Velizy-Villacoublay, France) and fabricated using a plastic micromachining method.

Microfeatures modelled in Solidworks were imported into SprutCAM (SprutCAM, Naberezhnye

Chelny, Russia), and converted into TAP files for milling on the CNC milling machine. Device

layers were fabricated with a Tormach PCNC 770 vertical milling machine (Tormach,

Waunakee, WI) from clear 2.0 mm thick sheets of PMMA. Devices were milled using 1/16”

(1.5875 mm) (Part # 01982156), 3/64″ (1.1906 mm) (Part # 07765431) and 0.020″ (508 μm)

endmills.

4.1.3 Plasma Treatment

The fabricated devices were then masked with scotch tape with the exception of the channel

walls and then subsequently treated with oxygen plasma in an oxygen plasma chamber (PVA

TePLA AG, Germany) at 60 W for 10 min.


4.1.4 Preparation of Solutions

Two solutions were prepared to obtain experimental data. 5 mL of 25% isopropyl alcohol (IPA)

solution (Sigma-Aldrich, St. Louis, Missouri, USA) in DI water was prepared and 4 drops of red

food colorant were added to the solution. 5 mL of deionized (DI) water with 4 drops of blue food

colorant was also prepared. The two solutions were then loaded in two separate 1 mL syringes

(BD Biosciences, San Jose, CA, USA).

4.1.5 Pressure-Driven Flow Experiment

The inlets of the PMMA device were connected to a syringe pump (Harvard Apparatus,

Southnatick,MA) by small-diameter tubing (PE-10, Warner Instruments, Hamden, CT).

Five-centimeter stainless steel 23-g blunt infusion cannulae were connected to the syringe. The

pump was set to flow at 30µL/min constant flow rate. Real time video of fluid flow was taken with

a Nikon D7200 DSLR Camera.

4.3 Results and Discussion

4.3.1 Contact Angle

The contact angles of two solutions, 25% IPA and DI water, on the PMMA device were

measured before and after oxygen plasma treatment for 10 min (Figure 4.3). Plasma treatment

time of 10 min was used to prevent hydrophobic recovery of PMMA for the duration of the

experiment. The contact angle of DI water dropped from 58.6o to 25.3o after plasma treatment

and for 25% IPA solution the contact angle dropped from 15 o to 4.8 o. Other fluidic properties

including surface tension and dynamic viscosity of these solutions were obtained from the

literature (Table 4.1) (J. Berthier et al. 2015) for use with the analytical models proposed. The

flow rate was converted to Δp using equation 4.5. See Appendix A 3.3 for derivation.


𝑝 = 𝑄𝐿

𝛼=

𝑄𝐿

ℎ𝑤3

8𝜇{1−

ℎ2

3𝑤2} (4.6)

Table 4.1 Physical properties of 25% IPA solution and DI water required for theoretical solutions.

25% IPA DI Water

Surface Tension, γ (mN/m), (Source: (J. Berthier et

al. 2015))

29 69.5

Dynamic Viscosity, µ (cP), (Source:(J. Berthier et

al. 2015))

2.4 1.2

Θ before plasma (deg), determined

experimentally

15 58.6

Θ after plasma (deg), determined experimentally 4.8 25.3

4.3.2 Pressure-Driven Flow Experimental Data for 25% IPA Solution and DI Water

Real-time videos of pressure driven flow through parallel wall channels with aspect ratio w/h > 1

were recorded for both 25% IPA solution (Figure 4.4) and DI water (Figure 4.5). The

penetration distance and velocity with respect to time were quantified for each solution using

ImageJ software (NIH). Results are presented in Table 4.2. For the same flow rate, 25% IPA

solution appeared to travel a longer distance compared to DI water. This is likely due to the low

Figure 4.3 Photographs of contact angles formed by DI water (blue) and 25% IPA in DI water (red) on a PMMA surface, before (A) and after (B) plasma treatment.

A B


contact angle (good wettability) of 25% IPA with PMMA compared to DI water with PMMA. Low

contact angle translates to high capillarity. This mean the flow due to the pressure difference

(flow rate) is assisted by this capillary force, resulting in faster travel distance of the fluid front.

Table 4.2 Experimental data for 25% IPA solution and water

Water 25% IPA

Time (s) Distance (m) Velocity (m/s) Distance (m) Velocity (m/s)

5 0.006 0.0012 0.006 0.0012

33 0.026 0.000788 0.024 0.000727

66 0.04 0.000606 0.051 0.000773

99 0.063 0.000636 0.073 0.000737

132 0.073 0.000553 0.09 0.000682

165 0.097 0.000588 0.107 0.000648

198 0.117 0.000591 0.13 0.000657

231 0.132 0.000571 0.151 0.000654

264 0.15 0.000568 0.172 0.000652

297 0.15 0.000505 0.183 0.000616

330 0.18 0.000545 0.197 0.000597


Figure 4.4 Individual frames from real-time video capture of pressure-driven suspended microflow between parallel walls for 25% IPA in DI water. A channel

with width (900µm), height (650µm) and length (20cm) devoid of a ceiling and a floor was used to simulate a high (width-to-height) aspect ratio parallel wall case. 25% IPA solution was dispensed at the inlet of the channel at a constant flowrate of 5.0 *10-10 m3/s. The solution travelled a total distance of 19.7cm in 330 sec.

A. t = 5 sec

Q. t = 198 sec

GG. t = 330 sec


Figure 4.5 Individual frames from real-time video capture of pressure-driven suspended microflow between parallel walls for water. A channel with width

(900µm), height (650µm) and length (20cm) devoid of a ceiling and a floor was used to simulate a high (width-to-height) aspect ratio parallel wall case. Water was dispensed at the inlet of the channel using a syringe pump at a constant flowrate of 5.0 *10-10 m3/s. The solution travelled a shorter distance compared to the 25% IPA solution, a total of 18cm in 330 sec.

A. t = 5 sec

B. t = 198 sec

C. t = 330 sec


4.3.3 Comparison between Theoretical and Experimental Pressure-Driven Flow

Data for 25% IPA Solution and DI Water

Analytical plots of penetration distance and velocity with respect to time were plotted using

equation 4.2 and 4.3. The experimental data collected from the experiments described above

were also plotted in the same graph. Figure 4.6 shows the plot for penetration distance with

respect to time. According to both the theoretical and experimental data, under same flow

conditions, the penetration distance for 25% IPA solution appears to be higher than DI water. As

briefly discussed before this is due to a lower contact angle obtained for 25% IPA solution

compared to DI water, both before and after plasma treatment, leading to a high capillary force

and resulting in a greater distance traveled. For both cases the experimental data appears to be

below the theoretical curve, suggesting that in general the fluid flows a short distance over a set

Figure 4.6 Penetration distance of the liquid front versus time for 25% IPA (orange curve), and water (blue curve) with a contact angle of 25.3o and 4.8o (after plasma-treatment) respectively on the wall surface. The dots are the

experimental results for 25% IPA (yellow dots) and water (purple dots). The experimental results are not a good fit with a square root law at the beginning of motion; progressively the experimental results begin to appear in close relation with the theoretical curve.


time. In addition, the experimental data does not follow a square root law near the beginning of

flow as the analytical curve predicts. Progressively with time, however, the experimental data

appears to be close in relation with the analytical curve.

Figure 4.7 shows the plot for velocity of the travelling fluid front with respect to time. According

to both the theoretical and experimental data, under same flow conditions, the velocity of the

fluid front for 25% IPA solution appears to be higher than DI water. Since the experimental

velocity measurements were made using penetration distance data (ie. velocity = (displacement

/ set time)) the same relationship between experimental data and theoretical curve is observed

as Figure 4.6.

Several environmental factors could account for the discrepancy between theoretical and

experimental data. First, the effects or wall surface roughness on fluid flow is not taken into

consideration in the analytical solutions. We know that surface roughness impacts contact

Figure 4.7 Velocity of 25% IPA (orange curve), and water (blue curve) versus time, with a contact angle of 25.3o and 4.8o respectively on the wall surface. The

dots are the experimental results for 25% IPA (yellow) and water (purple). The experimental data seems to suggest that the velocity was relatively constant after the first 50 seconds of fluid motion.


angles of liquid against a surface. Since the PMMA devices were micromilled, it is possible that

residual PMMA pieces attached to the walls of the device pinned the fluid to the wall preventing

it from moving forward until the fluid build-up at the fluid-front from the applied pressure force at

the inlet was large enough to overcome the opposing forces from the fluid being pinned to the

walls. Further studies may need to be conducted to assess the surface roughness of the PMMA

walls after milling to support this hypothesis. Second, the solutions used for this study are

volatile fluids, especially the 25% IPA solution. The analytical solution doesn’t take into account

the effects of evaporation and for microscale volumes the effects of evaporation are critical.

Therefore, it is possible that due to evaporation or loss of volume of fluid the fluid front, in our

experiments, were always behind the projected theoretical values. Lastly, the analytical solution

assumes that the cross-sectional area of the fluid front remains constant throughout the

experiment. In reality however, because there are two free interfaces (due to no ceiling and floor

condition) the ALIs are free to change. This may occur if the inertial forces due to fluid buildup is

significantly higher than the viscous forces of the fluid at any point in time. When this happens

the ALI increases in size which is not accounted for in the analytical solutions. More specifically

this affects the theoretical capillary force calculation, particularly the term that accounts for

cohesive forces between fluid molecules that oppose the adhesive forces between the fluid and

solid surface molecules (see Appendix A2).

In conclusion, although our proposed analytical solutions do not conform well with experimental

data, it follows a similar trend. In the future, further modifications can be made to the analytical

solution to account for factors mentioned above, namely surface roughness, evaporation effects

and changes in cross-sectional perimeter of the ALI.

51

5.0 Preliminary Assessment of the Lung Airway

Device

Although the physical design parameters and microfabrication procedure of the proposed airway

microfluidic device had been decided on in previous chapters, the final tissue engineering

design of the proposed in vitro model will require further investigation and empirical research.

The goal from a biological perspective is to mimic important features of the airway in vivo

microenvironment such as the tissue-tissue interface, tissue architecture and other tissue

properties, as well as the physicochemical microenvironment. Our plan is to co-culture a

monolayer of Calu-3 epithelial cells and HBSMCs on opposing sides of a suspended ECM

hydrogel inside the proposed PMMA microfluidic device to mimic the tissue organization and

tissue-tissue interface (ie. bronchial epithelium + connective tissue + smooth muscle) found in

the bronchioles of the respiratory tract. In addition, we aim to recreate similar tissue morphology

and tissue barrier properties of the bronchial epithelium.

Given that co-culturing of both Calu-3 cells and HBSMCs will depend on the attachment of both

cell types to the matrix, it is important to examine cell adhesion behavior of these cells to find an

optimal ECM hydrogel or mix. Furthermore, the ability of both cell types to not only attach but to

spread and proliferate will also need to be the subject of investigation for future goals of

5.0 Preliminary Assessment of the Lung Airway Device 52

developing a robust in vitro model that can sustain long-term co-culture and be used to study

airway mechanisms and responses.

Investigation of cell culture parameters that would produce similar tissue characteristics found in

the in vivo state is a critical step in the early stages of model development to assess its potential

as an in vitro model.

One of the crucial components of the lung airway tissue is the respiratory epithelium, which in

the bronchiolar region is composed of a cuboidal and ciliated monolayer of epithelial cells. The

Calu-3 cell line is often used as a respiratory epithelial cell model to study human respiratory

processes and diseases. Although numerous reports demonstrate the formation of Calu-3

monolayers on various forms of 2D ECM coated support (Zhu, Chidekel, and Shaffer 2010), our

system features the formation of Calu-3 monolayers on a 3D, purely ECM hydrogel that is

suspended inside a PMMA microfluidic device. Aside from the proper selection of protein

composition in the suspended ECM hydrogel, culture parameters, such as culture time and cell

seeding densities may have a major impact on the formation of this type of monolayer and

therefore will also need to be examined.

Calu-3 cultures in conventional culture systems, (eg. culture at an ALI on a permeable

membrane), and for a few cases in microfluidic systems, have been able to reproduce many

characteristic properties of the in vivo bronchial epithelium, such as a tight monolayer with good

barrier integrity and permeability properties, (Y. Shen et al. 2015), (B. Q. Shen et al. 1994),

(Foster et al. 2000a). One of the characterization methods often used to demonstrate tight

monolayer and good barrier integrity include staining for the presence of junction proteins

involved in forming tight cell-cell junctions (eg. intercellular and tight junctions). To determine

whether Calu-3 cultures in our system are able to reproduce some of these tissues properties,

we will need to examine the presence of these proteins in our culture.


HBSMCs have been shown to play critical roles including in airway responsiveness to irritants,

regulating airway tone and have demonstrated phenotypes typically seen in many lung

disorders such as asthma when conditioned to stimulation in culture (Shubber et al. 2015),

(Lambert et al. 1993),(Ebina et al. 1993). Thus, when developing airway in vitro models to study

airway responsiveness and to elucidate mechanisms of airway diseases, it is important to take

into consideration the influence of smooth muscle tissue. Within the lung, smooth muscle cells

exist in bundles of aligned population wrapped around the bronchial tubes in a helical fashion.

To recreate this type of morphology in our system, we will need to examine cell culture

parameters that will produce this type of smooth muscle cell morphology. Actin, a major

structural and functional cytoskeletal protein, is known to mediate diverse biological processes

like controlling cell shape, polarity, motility, contraction as well as cytokinesis. These processes

have a central involvement in airway responses for both healthy and diseased state bronchioles

(Panettieri et al. 1989) and actin expression is commonly used to detect changes in these

biological processes. It can also be used to visualize the cytoskeletal structure and morphology

of cells.

In this chapter, we conduct several studies to optimize culture parameters of our in vitro airway

model and perform preliminary assessment of the monocultures and co-cultures developed in

our system.

We conduct a study to select an optimal ECM hydrogel that would promote cell adhesion and

proliferation of both Calu-3 cells and HBSMCs. ECM protein(s) used for this study included,

Type I collagen (Col-1) and Matrigel, based on their abundance in most connective tissues and

basement membranes, respectively. Col-1 is a protein that is abundantly available within the

interstitial ECM, which surround most cells and provides structural support for tissues (Bonnans,

Chou, and Werb 2014). The basement membrane, a specialized form of ECM separating the

epithelium from underlying tissue, and its protein constituents are also known to play major roles

in epithelial cell organization and differentiation (Bonnans, Chou, and Werb 2014). As


mentioned before, these proteins also have the ability to stimulate complex cell behavior that

are difficult to replicate using synthetic matrices. We investigated cell adhesion behavior on

Col-1 and Matrigel alone as well as a mix of these protein(s) to expand our search for an optimal

ECM hydrogel.

A second study is conducted to determine the appropriate culture parameters including time and

cell seeding densities that would produce a confluent monolayer epithelium. Calu-3 cells are

cultured at different cell seeding density on a suspended ECM hydrogel for up to 21 days in our

microfluidic system to determine these culture parameters. We also determine HBSMC culture

parameters required to reach confluence when cells are cultured on the underside of a

suspended ECM hydrogel. HBSMCs were cultured independently for up to 14 days for this

study. As preliminary assessment of co-culture in our system, we examine one case where

Calu-3 cells are cultured independently, for 14 days and subsequently co-cultured with

HBSMCs for 7 days summing to a total 21-day culture period.

Furthermore, we look at whether tight Calu-3 monolayers are formed when cultured on a

suspended ECM hydrogel in our device. In particular we stain for the presence of F-actin and

zonula occludens-1 (ZO-1) proteins involved in the formation of tight cell-cell junctions (Wan et

al. 2000). We test this for both our monoculture and co-culture systems of Calu-3 cells. We also

examine the expression of F-actin and smooth muscle cell phenotypic marker, α-smooth muscle

actin (α-SMA), in both monocultures and co-cultures of HBSMCs in our systems. Actin

expression is also used to evaluated the affect of culture time on HBSMC alignment in our

systems.


Unless otherwise mentioned, device design parameters and fabrication method used for all

studies in this chapter were the same as described in Chapter 3.


5.1.1 Device Design and Fabrication

The device design for cell adhesion study was different than all other studies conducted. Device

layers were fabricated with a Tormach PCNC 770 vertical milling machine from clear 1.5 mm

thick PMMA sheets with dimensions, 100 mm X 25 mm. Devices were milled using 1/16”

(1.5875 mm) and 3/64″ (1.1906 mm) endmills.

The top and bottom layer

The top layer consisted of a series of rectangular channels, with curved edges and

through-holes for the inlets and outlets. Each channel had two through-holes, an inlet and outlet,

located inside at each short edge of the channel (Figure 5.1A). The channels were 1.25 mm tall,

4 mm in diameter and spanned 8 mm from the centre of the two through-holes located inside the

edges of the channels. The inlet and outlet through-holes were 2.75 mm in diameter. The

bottom layer consisted of channels in the same orientation and dimensions as the top layer. The

channel depths were adjusted to 0.7 mm. A second set of smaller rectangular channel with

curved edges were inserted inside each larger rectangular channels. The smaller channels were

0.5 mm deep, 2.0 mm wide and 4 mm in length. The inlet edges of the smaller channels were

mirrored with the inlet through-holes, located inside the edges, of the top layer Figure 5.1B.

PMMA layers were bonded using method described in Chapter 3.

5.1.2 Cells and Cell Culture

All reagents were purchased from Life Technologies (Carlsbad, CA, USA) unless otherwise

stated.

Calu- 3 cells were obtained from American Type Culture Collection (ATCC, Manassas, VA,

USA) and maintained in MEM- alpha (no nucleosides) media with 10% standard fetal bovine


serum (FBS), 1 % HEPES buffer (1M) and 1% streptomycin—penicillin. HBSMCs were obtained

from American Type Culture Collection (ATCC, Manassas, VA, USA) and maintained in SmBM

Basal Medium containing SingleQuot supplements (Lonza, Allendale, NJ, USA) and 1%

streptomycin—penicillin. Both cell types were maintained in regular tissue culture flasks in an

incubator at 37oC with 5% CO2. All experiments reported were conducted using cells between

passage 11 to 14 for Calu-3 cells and between 9 to 12 for HBSMCs.

The following steps were conducted in a class II, Type A2 biological safety cabinet (BSC).

5.1.3 Experimental Preparation

Previously prepared PMMA devices were disinfected by pipetting and aspirating 70% ethanol in

DI water through the compartments of the device two times. During a third rinse the 70% ethanol

in DI water solution was left in the device for 20 min and subsequently rinsed with phosphate

buffered saline (PBS), three times. After aspirating residual PBS solution from the device, it was

left inside the BSC for 20min to evaporate remaining PBS volatiles. This is a crucial step to

ensure proper loading of ECM hydrogels and selective coating of channel walls inside the

device. See Appendix A5 for a more detailed protocol.

Figure 5.1 Two layer PMMA microfluidic device for cell adhesion study. A) Top PMMA layer consisting of an array of 16 channels with an inlet and outlet ports in each channel. B) Bottom PMMA layer consisting of an array of 16 channels with smaller channel embedded within larger channels.

A

B


5.1.4 Cell Adhesion on Hydrogel Study

ECM preparation

Four different hydrogels or hydrogel composites were tested, (Table 5.1). Type 1 collagen and

Matrigel exclusively were two of the hydrogels and the others were hydrogel composites

including a mix of Col-1 and Matrigel. An example is presented in which the total hydrogel

volume is either 150µL or 157µL.

A 150µL collagen hydrogel solution was prepared by adding 125µL of Type I collagen (Col-I)

(rat tail, BD Biosciences, Bedford, MA, USA), at 10.31 mg/mL to 25µL of a basic solution (0.5 N

NaOH) to neutralize the acidic Col-I solution. For a final concentration of 6 mg/mL (Col-I) and 3

mg/mL total protein concentration of Matrigel, a mixture consisting of 87.5 µL of (Col-I) and 17.5

µL of 0.5 N NaOH was prepared and kept at 4oC for 1 hour (Sung et al. 2010). Matrigel was

thawed at 4oC for 1-hour solution and 52 µL was added to the mixture to make a total hydrogel

volume of 157 µL. Another hydrogel solution was prepared that consisted of a final

concentration of 3 mg/mL (Col-I) and 6 mg/mL total protein concentration of Matrigel. Similar to

the mixture prepared above, 45 µL of (Col-I) and 9 µL of 0.5 N NaOH was prepared and kept at

4oC for 1 hour before adding 103 µL of thawed Matrigel to the mixture. A 150µL of hydrogel

solution of Matrigel (BD Biosciences, Bedford, MA, USA) at 9 mg/mL total protein concentration

was also prepared. All protein bulk containers and prepared solutions were kept on icepacks

during and after preparation to prevent gelling before dispensing into device and subsequent

incubation period.

Hydrogel Loading

Prior to hydrogel loading the smaller channels of the middle PMMA layer were coated with 100

μg/mL bovine plasma fibronectin (FN) (Sigma, St. Louis, Missouri, USA) to facilitate adhesion of

the hydrogel to the channel walls. This was done by dispensing 9 μL of FN solution into the

smaller channels and incubating at room temperature for 20 min. After incubation the FN


Table 5. 1 Compositions ECM hydrogels tested for cell adhesion study.

Col-1[µL] Col-1 (6 mg/mL)

& Matrigel

(3mg/mL), [µL]

Matrigel

(6mg/mL) &

Col-1 (3 mg/mL),

[µL]

Matrigel [µL]

Col-1 (10.31

mg/mL)

125 87.5 45 0

0.5 N NaOH 25 17.5 9 0

Matrigel (9

mg/mL)

0 52 103 150

Total Volume 150 157 157 150

solution was aspirated out and the device was cooled down to prevent partial polymerization

during hydrogel loading. To do this the device was contained in a sealed Omnitray and kept at

-20 oC for 15 min. After cooling, the device was taken out of the Omnitray inside the BSC. To

position the device for hydrogel loading, the device was tilted at a 45o angle towards the

operator in an orientation such that the inlets ports were above the outlet. The previously

prepared hydrogel solutions were dispensed into the smaller channels of the middle PMMA

layer, one by one in a slow manner. The dispensing speed was adjusted depending on the

speed at which the hydrogel solutions wet the walls of the channel via surface tension. After gel

loading, the devices were incubated (37 °C, 5% CO2, for 2 hours) in a fully humidified chamber

for polymerization. Supplemented media used for Calu-3 cell maintenance was dispensed in

both the top and bottom compartments of each system in the device and incubated at (37 °C,

5% CO2, for 3-6 hours) to return any moisture that may have evaporated from the hydrogel

during polymerization.

Cell Culture

To investigate cell adhesion on the hydrogel solutions prepared, four devices were fabricated as

previously described, two for each cell type. For each cell type, one device was used for a 2-day

culture and the other for a 7-day culture. Each device had 16 systems, where 4 systems were


used for each hydrogel solution. Calu-3 cells and HBSMCs were trypsinized and suspended at

3000cells/μL and 500cells/μL, respectively, in their respective supplemented media. In order to

allow the cells to attach, 80 μL cell solutions were seeded into each system of all devices and

incubated at (37 °C, 5% CO2, for 12 hours) before flushing the channel with fresh media two

times to remove unattached cells. After initial flushing, the media was replaced every 24 h to

replenish nutrients in the media.

Cell Staining and Fixation

Cells were monitored for cell adhesion and viability by Calcein AM/ ethidium homodimer -1 (Life

Technologies, Carlsbad, CA, USA) and Hoechst 33342 (Life Technologies, Carlsbad, CA, USA)

nuclear stain on Day 2 & 7. Cells within each system were flushed with serum free media (Life

Technologies, Carlsbad, CA, USA) 3 times to replace residual solutions and incubated (37 °C,

5% CO2, for 12 hours) with 2μM Calcein AM, 4 μM ethidium homodimer -1 and Hoechst 33342

(Life Technologies, Carlsbad, CA, USA,) at (1:1000) dilution in serum-free media. Each volume

replacement for each channel was 80µL. After staining, cells were fixed with 4%

paraformaldehyde (PFA) (Sigma-Aldrich, St. Louis, Missouri, USA) and subsequently washed

with sterile PBS 3 times for imaging.

Microscopy and Image Analysis

Images were taken with EVOS FL Auto Cell Imaging System inverted microscope (Life

Technologies, Carlsbad, CA, USA). Devices did not need to be sterile during imaging

acquisition.

In order to quantify Calu-3 cell adhesion we analyzed the area coverage. This is because Calu-3

cells in culture forms networked islands, whether that be in culture flasks or on ECM hydrogels

(Figure 5.2). Given our objective of obtaining a confluent continuous monolayer of Calu-3

epithelium, area coverage is the most direct method of quantification for cell adhesion that is

relevant to our goal, as opposed to cell counts averaged over a certain surface area. The latter


approach introduces errors as it does not accurately account for the regions of the surface not

covered by Calu-3 cells. Hoechst nuclear stained images were analyzed for cell adhesion and

quantified with ImageJ software (NIH). Four systems were analyzed for each condition.

HBSMCs in culture tend to disperse uniformly across a surface, thus, for HBSMC cell adhesion

analysis we decided to use counts over a set surface area as a method of quantification.

Hoechst nuclear stained images were analyzed for cell adhesion and quantified with ImageJ

software (NIH). Four systems were analyzed for each condition

5.1.5 Long-term Cell Culture in Microfluidic Device

In all experiments, media replacement after initial wash was done every 24 hours. Each device

was kept in a separate sterile 128 mm × 86 mm Omnitray and Omnitrays were further kept in a

245 mm X 245 mm bioassay dish. For cultures lasting longer than 5 days, Omnitrays and

bioassay dishes were washed with soap, rinsed with water and sterilized with 70% ethanol in DI

water every 5 days of culture period to reduce the risk of contamination between Omnitrays,

bioassay dishes and our device.

Figure 5.2 Calu-3 cells cultured in a T-25 flask form networked islands. Scale bar = 400µm.

Figure 5.3 Sample preparation for microscopy of immunolabeling experiments. A) Microfluidic device with monocultures of Calu-3 cells and HBSMCs, post immunolabeling and fluorescent staining with a single sided blade lodged between the top and middle layer of the device. B) After detaching the top layer from the middle and bottom layer of the device the hydrogels were detached from the side walls of the channels using the sharp tip of a scalpel. C) The samples were then placed on a microscope slide. D) A single system with Calu-3 cells cultured and grown to confluence and placed on a glass microscope slide for imaging, as shown Hoechst stained image (inset). E) A single system with HBSMCs cultured and grown to confluence and placed on a glass microscope slide for imaging, as shown Phalloidin and Hoechst stained image (inset)Figure 5.2 Calu-3 cells cultured in a T-25 flask form networked islands. Scale bar = 400µm.

Figure 5.3 Sample preparation for microscopy of immunolabeling experiments. A) Microfluidic device with monocultures of Calu-3 cells and HBSMCs, post immunolabeling and fluorescent staining with a single sided blade lodged between the top and middle layer of


5.1.6 Cell Seeding Density Study

Calu-3 cell seeding and culture

ECM hydrogels were suspended, incubated and rehydrated as previously described. To

conduct cell seeding density study, cells were seeded at 5000, 4000, 3000, 1500 and 1000

cells/μL in supplemented media. In order to allow the cells to attach, 80 μL cell solutions were

seeded into each system of all devices and incubated at (37 °C, 5% CO2, for 12 hours) to allow

the cells to attach before flushing the channel with fresh media 2 times to remove unattached

cells. After initial flushing, the media was replaced every 24 h to replenish nutrients in the media.

Systems with cell seeding densities of 5000 and 4000 cells/µL were stained, fixed and imaged

on Day-2. The remaining systems were stained, fixed and imaged on day 3, 5, 8, 14 and 21.

HBSMC seeding and culture

ECM hydrogels were suspended, incubated and rehydrated as previously described. Prior to

cell loading, two 5mm thick,1cm X 0.5cm, sterile PDMS posts were placed in a sterile Omnitray

approximately 70 cm apart. HBSMCs were seeded at 500, 750 and 1000 cells/μL in its

respective supplemented media. 50μL cell solution was dispensed into the bottom channel. The

device was turned over and placed faced down on the two PDMS posts device and incubated

(37 °C, 5% CO2, for 12 hours) to allow the cells to attach. It is important to ensure the thickness

of the PDMS posts are the same to avoid tilting the surface of the ECM hydrogel onto which the

HBSMCs will adhere and grow. After 12 hours, the device was turned over one more time, to

have the HBSMC face down, and the channel was flushed with fresh media 2 times to remove

unattached cells. After initial flushing, the device was kept with the HBSMCs facing down for the

remainder of culture period. The media was replaced every 24 h to replenish nutrients in the

media. Systems were stained, fixed and imaged on day-7.



Cells were monitored for viability by a LIVE/DEAD assay (Calcein AM/ ethidium homodimer -1)

and Hoechst 33342 nuclear stain. Cells within each system were flushed with serum free media

3 times to replace residual solutions and incubated (37 °C, 5% CO2, for 30 min) with 2μM

Calcein AM, 4 μM ethidium homodimer -1 and Hoechst 33342 (1:1000 dilution in serum free

media). Each volume replacement for each channel was 88µL. After staining, cells were fixed

with 4% paraformaldehyde (PFA) and subsequently washed with sterile PBS 3 times for

imaging.

Imaging

Images were obtained with EVOS FL Auto Cell Imaging System inverted microscope. Devices

did not need to be sterile during imaging acquisition.

5.1.7 Calu-3 epithelium barrier structure

To ensure cells were forming cell junctions, Calu-3 cells were cultured (as previously described

in Calu-3 cell seeding and culture) at 3000 cells/µL seeding density for 21 days. Samples were

stained on day 7, 14 and 21 inside the microfluidic device. At the end of each culture time period

cells were fixed with 4% PFA and stained. Briefly, samples were blocked with blocking buffer

(3% bovine serum albumin (BSA) with 0.1% Tween 20 in PBS for 3 h at 37 °C; Sigma, St. Louis,

Missouri, USA) and immunolabeled with Rabbit anti-ZO-1 pAb (1:200 dilutions in blocking buffer

for 48 h at 4oC; Life Technologies, Carlsbad, CA, USA). After labelling samples were incubated

(37 °C, 5% CO2 for 2 hours) with AlexaFluor 568 goat anti-rabbit IgG (1:200 dilution in blocking

buffer; Life Technologies, Carlsbad, CA, USA) for conjugation with ZO-1, ActinGreen 488 (2

drops per ml of solution; Life Technologies, Carlsbad, CA, USA) to stain F-actin and Hoechst

33342 (1:1000 dilution) to stain the nuclei.


5.1.8 Actin labelling in HBSMCs

To ensure HBSMCs were expressing F-actin and phenotypic marker, α-SMA, cells were

cultured, as previously described (in HBSMC cell seeding and culture) at 500 cells/µL seeding

density for 14 days. Cells cultured on the underside of the suspended hydrogels were fixed with

4% PFA, permeabilized with 0.1% Triton X-100 (Sigma Aldrich, St. Louis, Missouri, USA) and

stained with Alexa Fluor 488 phalloidin (Life Technologies, Carlsbad, CA, USA) and with mouse

anti-α-SMA mAb (Sigma, St. Louis, Missouri, USA) to identify F-actin and α-SMA (Clone 1A4)

respectively. Briefly, samples were blocked with blocking buffer (3% BSA with 0.1% Tween 20 in

PBS for 3 h at 37 °C) and labelled with anti-α- SMA mAb (1:200 dilution in blocking buffer at 4oC

for 48 hours). After labelling, samples were incubated (37 °C, 5% CO2) with AlexaFluor 568 goat

anti-mouse IgG (1:200 dilution in blocking buffer; Life Technologies, Carlsbad, CA, USA) for

conjugation with α-SMA, Alexa Fluor 488 phalloidin (2.5% (v/v) in blocking buffer; Life

Technologies, Carlsbad, CA, USA) to stain F-actin and Hoechst 33342 (1:1000) to stain the

nuclei, in the device for 2 hours.

5.1.9 Co-culturing Calu-3 cells and HBSMCs

For co-culture experiments, ECM hydrogels were suspended, incubated and rehydrated as

previously described. Calu-3 cells were seeded at 3000 cells/µL and cultured, as previously

described, for 14 days. On day 14, prior to HBSMC loading, two 5 mm thick,1cm X 0.5cm, sterile

PDMS posts were placed in a sterile Omnitray approximately 70 cm apart. HBSMCs were

seeded at 500 cells/µL into the bottom channel and as described before, the device was turned

over to be rested on the two PDMS posts. The device was then incubated (37 °C, 5% CO2 for 12

hours) for cell attachment and turned over a second time for initial media replacement and

subsequent culture. The device was then incubated (37 °C, 5% CO2 for 7 days) with daily media

replacements.


5.1.10 Immunostaining Co-culture Samples

To ensure Calu-3 cells retained cell-cell junctions, already developed by day 14 of monoculture,

for the 7 days of co-culture and HBSMCs are expressing F-actin and α-SMA, samples were

stained on day 4 and 7. At the end of each culture time period, cells were fixed with 4% PFA,

permeabilized with 0.1% Triton X-100 and stained.

Briefly, samples were blocked with blocking buffer (3% BSA with 0.1% Tween 20 in PBS for 3 h

at 37 °C) and co-immunolabeled (4oC for 48 hours) with Rabbit anti-ZO-1 pAb (1:200 dilutions in

blocking buffer) and with mouse anti-α-SMA mAb (1:200 dilution in blocking buffer). After

labelling, samples were incubated (37 °C, 5% CO2, for 2 hours) with AlexaFluor 488 goat

anti-rabbit IgG (1:200 dilution in blocking buffer) for conjugation with ZO-1, AlexaFluor 568 goat

anti-mouse IgG (1:200 dilution in blocking buffer) for conjugation with α-SMA and Hoechst

33342 (1:1000 dilution) to stain the nuclei. Samples were subsequently washed with PBS to

remove excess fluorescent labelling solution.

Other co-culture samples (after fixation and permeabilization) were blocked with blocking buffer

(3% BSA with 0.1% Tween 20 in PBS for 3 h at 37 °C) and fluorescently labeled (37 °C, 5%

CO2, for 2 hours) with Alexa Fluor 488 phalloidin to stain for F-actin in both Calu-3 monolayers

and HBSMCs. Samples were subsequently washed with PBS to remove excess fluorescent

labelling solution.

5.1.11 Microscopy of Immunolabeled Samples and Data Analysis

For imaging, excess fluorescent labelling solution was removed with PBS from each system and

samples were analyzed with a fluorescence microscope (EVOS FL Auto Cell Imaging System).

To obtain image clarity, immunolabeled samples were extracted from the device, mounted on a

1.2 mm thick 75 X 25 mm glass microscope slide with PBS and analyzed with a fluorescence

microscope (EVOS FL Auto Cell Imaging System).


Sample preparation

To extract samples from the device, the top layer of the PMMA microfluidic device was detached

from the bottom two layer using a single edge steel blade (Figure 5.3 A). The blade edge was

lodged between the top and the middle layer at one corner of the device. While the blade was

lodged in, it was moved along the four edges of the device to slowly open the PMMA-PMMA

bonds between the top and middle layer. The blade was then moved across the device while still

lodged between the PMMA layers to detach the top layer. Once the top layer was removed, the

tip of the sharp scalpel was used to detach the edges of the hydrogel (highlighted in red in

Figure 5.3 B) from the edge of the channel. The hydrogels were then gently rinsed using a

pipette with PBS until the hydrogels completely detached from the device and were placed on a

1.2 mm thick 75 X 25 mm glass microscope slide for imaging (Figure 5.3 C, D).

Data Analysis

Nuclear stained images were used for cell alignment data collection as the alignment angle of

the nuclei, in most cases, appeared to be parallel to the alignment angles of actin stress fibers

(Figure 5.4). ImageJ software, more specifically the plugin “Orientation J”, was used to

measured the angles of alignment.

Statistical Analysis

All values are presented as mean ± standard deviation. In total, 4 experiments (n=4) were

conducted, meaning results from four systems were used for each condition on each day for

each of the two different cell types. Single-variable analysis of variance (ANOVA) was used for

multiple comparisons within a study (effect of ECM hydrogel on cell adhesion of each cell type),

and multiple comparison tests were performed using Tukey’s method. Two-way ANOVA was

performed to analyze the effects of culture time and ECM hydrogel type on cell adhesion of each

cell types. Multiple comparison tests were conducted using Sidak’s method. For both types of

analysis, differences were considered significant for P < 0.05.


Figure 5.3 Sample preparation for microscopy of immunolabeling experiments. A) Microfluidic device with monocultures of Calu-3 cells and HBSMCs, post

immunolabeling and fluorescent staining with a single sided blade lodged between the top and middle layer of the device. B) After detaching the top layer from the middle and bottom layer of the device the hydrogels were detached from the side walls of the channels using the sharp tip of a scalpel. C) The samples were then placed on a microscope slide. D) A single system with Calu-3 cells cultured and grown to confluence and placed on a glass microscope slide for imaging, as shown Hoechst stained image (inset). E) A single system with HBSMCs cultured and grown to confluence and placed on a glass microscope slide for imaging, as shown Phalloidin and Hoechst stained image (inset).

A B

D E

C


5.2 Results

5.2.1 Cell Adhesion Study

We first investigated cell adhesion and growth of Calu-3 and HBSMCs on various ECM hydrogel

solutions. We developed a simple method where a hydrogel solution was loaded into a

non-suspended channel inside a microfluidic device and polymerized to form a 500 µm thick

hydrogel layer. Calu-3 cells and HBSMCs were seeded and cultured for 7 days. Cell adhesion

on four different hydrogel compositions were compared and analyzed on day 2 and 7 of culture

to find an optimal hydrogel solution for co-culture. For each hydrogel solutions, 4 replicate

systems were analysed (n = 4) for each cell type, resulting in 32 systems analyzed on each day.

At the end of culture period, cells were stained with CalceinAM and Hoechst for cell counting

Figure 5.4 Immunostaining of HBSMCs culture on the underside of an ECM hydrogel suspended in a microfluidic device for 7 days. Cells were immunolabled

with Alexa Fluor 488 phalloidin for F-actin in green. Hoechst 33342 was used to stain the nuclei in blue. Scale bar = 200µm.


and quantitative analysis of overall cell adhesion. Cell adhesion of Calu-3 cells were correlated

with percentage of area covered by attached cells to the ECM hydrogels.

Calu -3

We noticed that on day 2, Calu-3 cells formed networks with neighbouring cells establishing

partial monolayers on Col-1 and both hydrogel mixtures, [Col-1 (6µg/µL) & Matrigel (3 µg/µL)]

and [Col-1 (3µg/µL) & Matrigel (6 µg/µL)], covering large regions of the hydrogel surface area

(Figure 5.5 A – C). Cells displayed the least coverage for the hydrogel consisting only of

Matrigel (Figure 5.5 D). Interestingly, by day 7 of culture the cells detached from Col-1 and

[Col-1 (6µg/µL) & Matrigel (3 µg/µL)] surfaces, (Figure 5.5 E, F), while adhesion behavior on

[Col-1 (3µg/µL) & Matrigel (6 µg/µL)] remained unchanged (Figure 5.5 G). Adhesion behavior

on Matrigel appeared to have no notable changes either (Figure 5.5 H).

Morphological analysis of area covered by adhered Calu-3 cells on day 2 confirmed significant

statistical differences between area coverage on [Col-1 (6µg/µL) & Matrigel (3 µg/µL)] compared

to Matrigel (***P<0.001) and area coverage on [Col-1 (3µg/µL) & Matrigel (6 µg/µL)] compared

to Matrigel (****P<0.0001) (Figure 5.6 A). Statistical difference was also observed on area

coverage between Col-1 and [Col-1 (3µg/µL) & Matrigel (6 µg/µL)] along with area coverage

between Col-1 and Matrigel (**P< 0.01). No statically significant difference (P>0.05) was found

in area coverage between Col-1 and [Col-1 (6µg/µL) & Matrigel (3 µg/µL)] as well as between

[Col-1 (6µg/µL) & Matrigel (3 µg/µL)] and [Col-1 (3µg/µL) & Matrigel (6 µg/µL)] (P>0.05).

Analysis of area covered by adhered Calu-3 cells on day 7 indicated that only hydrogel

composite [Col-1 (3µg/µL) & Matrigel (6 µg/µL)] yielded an area coverage above 80% (Figure

5.6 B). Statistically significant differences were observed in area coverage [Col-1 (3µg/µL) &

Matrigel (6 µg/µL)] and all other compositions tested in this study (****P<0.0001). Statistical

difference was also observed on area coverage between Col-1 and Matrigel along with area

coverage between [Col-1 (6µg/µL) & Matrigel (3 µg/µL)] and Matrigel (***P< 0.001). No


significant difference (P>0.05) was found in area coverage between Col-1 and [Col-1 (6µg/µL) &

Matrigel (3 µg/µL)].

Interestingly, on day 7 of culture, cell adhesion on Col-I and [Col-1 (6µg/µL) & Matrigel (3 µg/µL)]

surfaces appeared to have decreased compared to day 2, indicating detachment of cell from

these hydrogel surfaces. Two-way ANOVA analysis confirmed statistical significance in the

change of area coverage between day 2 and 7 for Col-I and [Col-1 (6µg/µL) & Matrigel (3

µg/µL)]] surfaces. Sidak’s multiple comparison test was performed (****P<0.0001) (Figure 5.7).

No significant difference (P>0.05) was found in change of area coverage between day 2 and 7

for Matrigel and [Col-1 (3µg/µL) & Matrigel (6 µg/µL)]. Area coverage on [Col-1 (3µg/µL) &

Matrigel (6 µg/µL)] surface remained above 80% from day 2 through to 7 but nothing can be

concluded yet as cell adhesion of HBSMCs on these hydrogels must also be investigated. This

is to accommodate co-culture and proliferation of both Calu-3 and HBSMCs on a selected

hydrogel composition.

HBSMC

Similar to the setup used for Calu-3 cell adhesion study, HBSMCs were cultured on four

different 500 µm thick hydrogel layers inside a PMMA microfluidic device for 7 days and

analyzed. On Day-7 of culture, HBSMCs appeared to be uniformly dispersed (without

networks) for hydrogels Col-1, [Col-1 (3µg/µL) & Matrigel (6 µg/µL)] and [Col-1 (6µg/µL) &

Matrigel (3 µg/µL)] (Figure 5.8 A-C). For Matrigel, HBSMCs appeared to have formed

aggregates and attached in small regions of the hydrogel surface but did not spread evenly

across the surface, (Figure 5.8 D).


Figure 5.5 Calu-3 epithelial cell adhesion on ECM hydrogel in a PMMA microfluidic device. Calu-3 epithelial cells were cultured on 500µm thick ECM

hydrogels containing various protein mixtures in a PMMA microfluidic device for 7 days. Adhered cells were stained with Hoechst 33342 for the nuclei and observed under fluorescence microscopy on Day 2 and 7. A, E: Col-1 ECM hydrogel. B, F: ECM hydrogel misture of Col-1 at 6µg/µL and Matrigel at total protein concentration of 3 µg/µL. C, G: ECM hydrogel mixture of Col-1 at 3µg/µL and Matrigel at total protein concentration of 6 µg/µL. D, H: Matrigel ECM hydrogel. Area enclosed within dashed lines are hydrogel surfaces. Scale bar 400 µm. I: inset of A, scale bar = 200 µm.

Day 2 Day 7


Figure 5.6 Area covered by adhered Calu-3 cells (%) on ECM hydrogel, containing various compositions of protein, inside a PMMA airway microfluidic device. (A) Effect of hydrogel composition on Calu-3 cell adhesion on day

2. (B) Effect of hydrogel composition on Calu-3 cell adhesion on day 7. Data presented as mean ± SD (n = 4). Data were analyzed using one-way ANOVA followed by Tukey’s multiple comparison test. Brackets indicate statistical significance, (***P<0.001, ****P<0.0001). Some significant comparisons have not been shown for clarity.

A B

Figure 5.7 Comparison of area covered by adhered Calu-3 cells (%) on various ECM hydrogel compositions between day 2 and 7. Data presented as

mean ± SD (n = 4). Two-way ANOVA was performed followed by Sidak’s multiple comparison test to analyze the effects of hydrogel compositions and culture time on the area covered by adhered cells. Brackets indicate statistical significance, (****P<0.0001). n ± SD (n = 4).


We also examined HBSMC adhesion and growth behavior over a 7-day culture period to

determine an optimal hydrogel composition for HBSMC culture. Matrigel was not considered in

this study. On Day-2 of culture, cells appeared to be well dispersed on all three different

hydrogel compositions. No notable differences in number of attached cells were observed

between the hydrogel compositions tested (Figure 5.9 A-C). This was confirmed by quantitative

analysis, where no statistically significant differences were observed (P>0.05) in cells attached

per mm2 of hydrogel between the three hydrogel compositions test on Day-2 (Figure 5.10 A).

Similarly, on day 7 of culture, cells appeared to be more confluent and uniformly dispersed

(Figure 5.9 D-F). Figure 5.10 B confirms this observation as no statistically significant

differences were found between cells attached per mm2 between the hydrogels tested.

To investigate cell proliferation, we combined the results of HBSMCs attached on hydrogels for

day 2 and 7, (Figure 5.11). The graph shows an increase in overall cell number for all three

hydrogel compositions tested from day 2 to 7 indicating cell proliferation.

Since HBSMC adhesion and proliferation behavior appears to be relatively constant between

the three different hydrogels tested, the selection of hydrogel composition for co-culture of both

Calu-3 and HBSMCs will depend on the adhesion efficiency of Calu-3 cells on these hydrogel

compositions. We discovered from the results above that hydrogel composition [Col-1 (3 µg/µL)

& Matrigel (6 µg/µL)] yields cell area coverage above 80% after 7 days of culture, which is much

higher than all other hydrogels tested. Thus from the evidence provided above, hydrogel

composition [Col-1 (3 µg/µL) & Matrigel (6 µg/µL)] will be used for all subsequent studies in this

report.


Figure 5.9

Figure 5.8 Fluorescence image of HBSMCs on different hydrogel compositions cultured inside a PMMA microfluidic device for 7 days. Images

were obtained on day 7. Adhered cells were stained with Calcein AM for live cell imaging. A: Col-1 ECM hydrogel. B: ECM hydrogel mixture of Col-1 at 3µg/µL and Matrigel at total protein concentration of 6µg/µL. C: ECM hydrogel mixture of Col-1 at 6µg/µL and Matrigel at total protein concentration of 3µg/µL, scale bar = 500 µm. D: Matrigel ECM hydrogel, scale bar = 1000 µm. Area enclosed within dashed lines are hydrogel surfaces.

A

B

C

D


Day2 Day 7

Col-1

Col-1 (3µg/µL)

& Matrigel

(6µg/µL)

Col-1 (6µg/µL)

& Matrigel

(3µg/µLg/µL)

Figure 5.9 Fluorescence image of HBSMCs on different hydrogel compositions cultured inside a PMMA microfluidic device for 2 and 7 days. Images were

obtained on day 2 and 7. Adhered cells were stained with Calcein AM and Hoechst 33342 for live cell and nuclear staining. A, D: Col-1 ECM hydrogel. B, E: ECM hydrogel mixture of Col-1 at 3µg/µL and Matrigel at total protein concentration of 6µg/µL. C, F: ECM hydrogel mixture of Col-1 at 6µg/µL and Matrigel at total protein concentration of 3µg/µL. Scale bar = 400 µm.

A D

B E

C F


Figure 5.10 Number of adhered HBSMCs on various ECM hydrogels during 7-day culture period. The graphs represent the number of attached HBSMCs per mm2 of

hydrogel surface inside a PMMA microfluidic device on day 2 (A) and day 7 (B) of culture. Data presented as mean ± SD (n = 4). Data were analyzed using one-way ANOVA. No statistically significant differences in cells attached / mm2 observed between hydrogel compositions.

A B

Figure 5.11 Cell Adhesion and growth of HBSMCs cultured for 7 days on various ECM hydrogels. The graphs represent a comparison of the number of

attached HBSMCs per mm2 of hydrogel surface inside a PMMA microfluidic device on day 2 (A) and day 7 (B) of culture. Graph shows an increase in HBSMCs attached per mm2 between day 2 and 7. Data presented as mean ± SD (n = 4).


5.2.2 Calu-3 Monolayer Formation and seeding density

For this study, we noticed that Calu-3 cells reached confluence between 2 and 3 weeks, when

the gel was suspended inside the device, but it took a shorter time, about 7 to 9 days, when the

hydrogel was supported (not suspended) inside the PMMA microfluidic device. To obtain an

epithelial monolayer, it is important to reduce regions of the hydrogel surfaces that are not

covered by cells, as much as possible. Different cell seeding densities were tested in preliminary

trials on a suspended hydrogel, and it was determined that cell seeding densities above

3000cells/µL resulted in cells forming 3-D cell clusters that do not resemble an epithelial

monolayer and are difficult to wash away. Thus, three different cell seeding densities were

tested 1000 cells/µL, 1500 cells/µL and 3000 cells/µL to determine culture period that would

yield a monolayer epithelium with maximum hydrogel surface area coverage (Figure 5.12).

Fluorescence images of Calcein AM and Hoechst stained Calu-3 cells were obtained on Day 3,

5, 8, 14 and 21. As shown on Figure 5.12 G-I, it was found that by day 14 cell seeding density of

3000 cells/µL appeared to have produce a more consistent and repeatable monolayer

epithelium with the highest area coverage. By day 14 cell seeding density 1000 cells/µL had

large regions of hydrogel not covered by cells (Figure 5.12 C) while cell seeding density 1500

cells/µL had smaller empty regions and larger area coverage but results were more inconsistent

compared to 3000cellls/µL (Figure 5.12 D-F).

Hydrogel area coverage by Calu-3 cells were quantified and it was determined that by day 21

both 1500 cells/µL and 3000 cells/µL cell seeding densities produced epithelium monolayer with

area coverage above 90% and 1000 cells/µL produced area coverage above 60% (Figure

5.13). By day 3 of culture the maximum area coverage obtained was 60% by cell seeding

density 3000 cells/µL. 3000 cells/µL seeding density yielded a mean area coverage above 80%

by day 14 and formed more consistent and stable monolayers by day 21. Cell density 1500

cells/µL yielded a mean area coverage above 70 % by day 14 and 1000 cells/µL cell seeding

density produced above 40% coverage by day 14.


Figure 5.12 Area of ECM hydrogel covered by cultured Calu-3 epithelial inside a PMMA microfluidic device. Cells were cultured on a suspended ECM hydrogel for 14

days and stained with Calcein AM on day 3, 8 and 14. A-C: fluorescence micrographs of cells seeded at 1000 cells/µL on day 3, 8 and 14 of culture respectively. D-F: cell seeding at 1500 cells/µL. G-H: cell seeding at 3000 cells/µL. Area enclosed within dashed lines are hydrogel surfaces. Scale bar = 200 µm.

A B C

E F

G H I

D

Day 3 Day 8 Day 14

1000 cells/µL

1500

cells/µL

3000 cells/µL


Figure 5.13 Area covered by adhered Calu-3 cells (%) on a suspended ECM hydrogel, inside a PMMA airway microfluidic device, cultured over a 21-day period. The graph represents the area covered by attached Calu-3 cells on hydrogel

surface on days 3,5,8,14 &21 of culture for cell seeding density 1000cells/µL, 1500cells/µL and 3000 cells/µL. By day 21 the mean area coverage for seeding densities 1500cells/µL and 3000 cells/µL appears to have reach well above 90%. Data presented as mean ± SD (n = 4).

5.2.3 HBSMC Seeding Density Study

We found through preliminary analysis that HBSMCs seeded at 500cells/µL achieved full

confluence by day 7 when cultured on the underside of suspended hydrogel mixture [Col-1

(3µg/µL) & Matrigel (6 µg/µL)]. Immunostaining with Phalloidin (F-actin) and Hoechst (nucleus)

revealed that cells tended to primarily align in parallel with the inlet-outlet axis of the channel

(Figure 5.14 A). Further analysis was done to confirm this observation. Alignment of cells

appeared to vary widely near the edges of the hydrogel where the cells were in contact with both

PMMA and the hydrogel surface.


5.2.4 Monoculture Characterization

Calu -3 and HBSMCs were cultured separately on the topside and underside, respectively, of a

suspended hydrogel mixture previously selected. Calu-3 cells were cultured for 21 days and

stained with Actin Green 488 to label the actin cytoskeleton and F- actin, Rabbit anti-ZO-1 pAb

to label tight junction proteins and Hoechst to label the nuclei. Fluorescence images were

obtained on Day 7, 14 and 21 of culture (Figure 5.14 B). Similarly, HBSMCs were cultured for

14 days and stained with Alexa Fluor 488 phalloidin to label F-actin and mouse anti-α-SMA mAb

to label α-SMA on days 4, 7 and 14. HBSMC orientation was also quantified and analyzed for

one representative sample on each day the samples were stained.

O

I

A B

Figure 5.14 Immunostaining of Calu-3 and HBSMCs co-cultured on opposing sides of an ECM hydrogel suspended in a microfluidic device. Calu-3 cells were

seeded on Day-0 and HBSMCs were seeded on Day-14. Representative images of co-culture on day 7 (21-day total culture) are presented here. Cells were immunelabelled with Alexa Fluor 488 phalloidin for F-actin in green. Hoechst 33342 was used to stain the nuclei in blue. Images were taken at the same X-Y location but on different Z-planes. F-actin is expressed in both Calu-3 cells (A) and HBSMCs (B). Scale bar = 500µm.


Calu-3 Monoculture Characterization

Actin Green 488 staining of F-actin and actin cytoskeleton revealed the presence of continuous

rings of F-actin along the cell borders for all three days together with a meshwork of actin

stress-fibres in the cytoplasmic regions (Figure 5.15 A-C). Epithelial barrier integrity is

maintained through interactions occurring at the cell–cell and cell–matrix interface by focal

adhesion complexes including tight junctions (TJs), adherens junctions, desmosomes and gap

junctions. TJs are organized by the interactions between various transmembrane proteins,

intracellular plaque zonula occludens (ZO-1, ZO-2, ZO-3), other proteins and the actin

cytoskeleton (Grainger et al. 2006). In our system, ZO-1 pAb staining displayed cobblestone

morphology and well-defined polygonal rings localized to the periphery of the cells,

predominantly in Day 14 and Day 21 of culture (Figure 5.15 E, F). This type of morphology is

common in Calu-3 monolayers, documented widely in the literature, and is indicative of stable

barrier properties formed in monolayers. ZO-1 TJ proteins were not observed in samples

obtained in day 7 of culture (Figure 5.15 D), suggesting that stable monolayers were not formed

by this time. Rings of F-actin in the merged images (Figure 5.15 G-I) appeared to be just below

the ZO-1 TJ proteins located in the junctional zone.

HBSMC Monoculture Characterization

Actin cytoskeleton of smooth muscle cells is a dynamic structure that play critical roles in the

development of mechanical properties of smooth muscle tissue and regulation of the

development of mechanical tension (eg. during smooth muscle contraction). Actin as a

cytoskeletal protein has major functions in mediating diverse processes such as motility

contraction and cytokinesis. It also plays major structural functions in controlling cell shape and

polarity. To localize the actin cytoskeleton, we co-stained our cultured HBSMC samples with

Alexa Fluor 488 phalloidin to label F-actin and mouse anti-α-SMA mAb, to label α-SMA. Images

were taken at a location away from where the side walls of the channels would have been

before the samples were taken out of the device. F-actin stained immunofluorescence images


indicated the presence of actin stress fibres within the cytoplasmic structure of HBSMCs on all

three days (Figure 5.16 A-C). α-SMA, a phenotypic marker of smooth muscle cells, was also

observed in the same samples on all three days (Figure 5.16 D-F). Merged images (Figure

5.16 G-I) show the distinction between F-actin and α-SMA within the cell cytoplasm. An increase

in cell number is also observed from day 4 to day 14 of culture, as indicated by the nuclear stain,

Hoechst in blue. Actin stress fibres appeared to be predominately oriented in the direction

parallel the length of the channel. Quantitative measurement of angle of orientation was

performed to further investigate this observation.

HBSMC Angle of Orientation

F-actin and α-SMA co-stained HBSMCs appeared to be aligned in the longitudinal direction,

along the length of channel. To measure alignment angles, Hoechst stained nuclei were used as

the nuclei appeared to align with the orientation of the actin fibres (Figure 5.17). For this study,

images analyzed were taken at a location away from where the side walls would have been

before the samples were removed from the device. Total number of cells analyzed were kept

constant at n = 134. A frequency distribution of the alignment angles was plotted in polar graphs

for day 4, 7 and 14 of monoculture (Figure 5.18). Higher degree of alignment is correlated with a

narrow distribution of alignment angles, in other words higher frequency for a smaller number of

10-degree ranges.

On day 4, the frequency distribution of the alignment angles appeared to be widespread with 12

occurrences or more of alignment angles in three different 10 degree ranges, 700 to 800, 800 to

900 and 900 to 1000 (Figure 5.18 A). The highest frequency within the 3 ranges was just above

25 and the lowest was just above 15. For day 7 of culture, frequency value greater than 20 also

appeared for 3 different 10 degree ranges, 800 to 900, 900 to 1000 and 1000 to 1100, however a

greater discrepancy is observed between these ranges with the highest frequency being around


Day 7 Day 14 Day 21

A B C

D E F

G H I

Figure 5.15 Immunofluorescence staining of Calu-3 monolayers, cultured on a suspended ECM hydrogel inside a microfluidic device, at day 7, 14 and 21. Cells were stained with Actin Green 488 for actin cytoskeleton in

green(A-C) and Rabbit anti-ZO-1 pAb for tight junction proteins in red(D-F). Hoechst 33342 was used to stain the nuclei in blue. G- I Merged image of actin cytoskeleton, tight junction proteins and the nuclei. Scale bar = 20µm.


H

A B C

D E F

G I

Day 4 Day 7 Day 14

Figure 5.16 Immunostaining of HBSMC monocultures on the underside of a suspended hydrogel in a microfluidic device, at day 4, 7 and 14. Cells were immunelabelled with Alexa Fluor 488 phalloidin for F-actin in green

(A-C) and mouse anti-α-smooth muscle actin mAb for α-smooth muscle actin in red (D-F). Hoechst 33342 was used to stain the nuclei in blue. G- I Merged image of F-actin, α-smooth muscle actin and the nuclei. Scale bar = 100µm.


50 for the range 900 to 1000 and the lowest being just below 30 for the range 800 to 900 (Figure

5.18 B). A narrower distribution of alignment angles was found for HBSMCs on day 14 of

culture. As seen on (Figure 5.18 C) the nuclei of the cells on day 14 of culture appeared to be

oriented in the northerly direction with a slight tilt towards the east. Only two ranges, 600 to 700

and 700 to 800 appear to have a frequency greater than 20 (Figure 5.18 D). Between these two

ranges the highest frequency, greater than 50, was observed for the range 700 to 800 and the

lowest frequency, just above 30, was observed for the range 600 to 700. Thus, preliminary

analysis of alignment angles for one representative sample on day 4 of culture spanned from

slightly in the northwest direction to slightly in the northeast direction with a mean alignment

angle of 84o and standard deviation of 37o. For a day-7 culture sample, a mean alignment angle

of 96o and standard deviation of 14o was observed while for a day-14 sample a mean alignment

angle of 72o and standard deviation of 11o was observed. These observations were consistent

throughout replicate experiments for each day. See Appendix A4 for the distribution plots of

other replicate systems.

5.2.5 Co-culture Characterization

The next step in the preliminary assessment of this in vitro airway model was to co-culture both

Calu-3 and HBSMCs and characterize the cultures in a similar way the monocultures were

characterized. To do this, we cultured Calu-3 cells at seeding density of 3000 cells/µL for 2

weeks and seeded HBSMCs on the 14th day of culture at 500 cells/µL. The two cell types were

in co-culture for 7 days (total 21-day culture for Calu-3 cells and 7-day culture for HBSMC).

Samples were stained on day 4 and 7 of co-culture.

Rhodamine-phalloidin staining of F-actin revealed the presence of perijunctional rings of F-actin

in both cell lines, together with a fine meshwork of cyto-plasmic stress fibres orientated

perpendicular to the apicobasal axis. Cells in co-culture were stained with Alexa Fluor 488


Figure 5.18 Immunolabeled fluorescent images of HBSMC

Figure 5.17 Immunolabelled fluorescent images of HBSMC monoculture at day 14, culture on the underside of a suspended hydrogel in a microfluidic device. Cells were co-stained with Alexa Fluor 488 phalloidin for F-actin in green and mouse anti-α- smooth muscle actin mAb for α- SMA in red. Hoechst 33342 was used to stain the nuclei in blue. Scale bar = 400µm.

A B

D C

Figure 5.18 Polar graphs showing frequency distribution of the angle of HBSMC alignment with respect to the inlet-outlet axis of the microfluidic channel. HBSMCs were culture on the underside of an ECM hydrogel suspended inside a

microfluidic device for 14 days. Radial (horizontal) axis represent the frequency of angle of alignment. Each bars represent a 100 range. Arrows point to the outlet of the microfluidic channel. Polar graphs represent frequency distribution for one representative sample on Day 4 (A), Day 7 (B) and Day 14 (D) of culture (n = 134). (C) Hoechst (nuclei) stained sample on Day 14 of culture, scale bar = 200µm. Mean alignment angles and standard deviations of each system are indicated on the top right corner of each graph for each system. A general trend of narrower distribution is observed with increasing culture time.

Mean = 84° SD = 37°

Mean = 96° SD = 14°

°

Mean = 72° SD = 11°


phalloidin to label F-actin and Hoechst to label the nuclei of both cell types after a 7-day

co-culture period. Fluorescence micrographs revealed the presence of densely packed actin

filaments along the periphery of the Calu-3 monolayer as seen in monoculture characterization

of Calu-3 monolayers (Figure 5.19 A). This morphology was observed uniformly across the

monolayer formed by Calu-3 cells in culture with HBSMCs. The fluorescence images also

showed the presence of actin stress fibers oriented along the longitudinal direction, parallel to

the length of the channel, on the HBSMC culture side similar to the morphology seen in

monoculture characterization (Figure 5.19 B).

Further co-culture characterization was performed to highlight phenotypic differences between

the two cell types and to determine if the characteristic properties of Calu-3 monolayer and

HBSMCs, as observed in monocultures inside our proposed micrfluidic device, can be

reproduced in the co-culture systems.

Calu-3 Monolayer Characterization in Co-culture with HBSMCs

Co-culture samples were co-stained for α-SMA (red) and ZO-1 TJ proteins (green) and showed

no double positive cells (Figure 5.21). However, since Hoechst is a nuclear stain for both cell

types, images obtained of co-culture samples often showed the nuclei of both cell types. This

problem was less prominent since the two cell types were co-cultured on opposing sides of a

650µm thick hydrogel and the images were taken on different planes along the cross-section of

the hydrogel.

Calu-3 monolayers expressed cobblestone morphology and well-defined polygonal rings

localized to the periphery of the cells both on day 4 and 7 of co-culture (Figure 5.21 C, D),

similar to the morphology observed in monoculture characterization. We know from our

monoculture characterization of Calu-3 monolayers that ZO-1 expression was present on day

14 of culture. Since HBSMCs were seeded on day 14 of Calu-3 culture, the expression of ZO-1


on day 4 and 7 of co-culture suggests that co-culture conditions did not have a notable effect on

the tight junctions formed by day 14 of Calu-3 culture.

HBSMC Characterization in Co-culture with Calu-3 cells

α-SMA expression was present in HBSMCs co-cultured with Calu-3 cells on both day 4 and 7

(Figure 5.21 A, B). The morphology appears to be very similar to the morphology observed in

monoculture characterization of HBSMCs, suggesting that co-culture conditions did not have a

notable effect on the culture. As seen before in monoculture characterizations, actin fibers

appeared to be aligned in the longitudinal direction, along the length of channel.

Actin fiber orientation angles were measured as before and the frequency distribution was

plotted in polar graphs (Figure 5.20). Total number of cells analyzed were 134. On Day 4 of

co-culture the frequency distribution of the alignment angles appeared to be widespread with 12

occurrences or more of alignment angles in 5 different 10 degree ranges, 200 to 300, 500 to 600,

700 to 800 and 800 to 900 (Figure 5.20 A). The highest frequency within the 5 ranges was just

above 16 for the range 800 to 900 and the lowest was at 12. For Day 7 of co-culture, frequency

value greater than 20 appeared for 3 different 10 degree ranges, 600 to700, 700 to 800 and 800 to

900. However, a greater discrepancy is observed between these ranges with the highest

frequency being around 45 for the range 700 to 800 and the lowest being just below 30 for the

range 800 to 900 Figure 5.20 B. A narrower distribution of alignment angles of HBSMCs were

observed for day 7 of co-culture. The angle of orientation spanned from the northwest direction

to the northeast direction with a mean alignment angle of 78o and standard deviation of 43o was

observed for our representative sample of day 4 co-culture. For a day-7 sample, a mean

alignment angle of 73o and standard deviation of 21o was observed. These observations were

consistent throughout replicate experiments for each day. See Appendix A4 for the distribution

plots of other replicate systems.



seeded on Day 0 and HBSMCs were seeded on Day 14. Representative images of co-culture on Day 7 (21-day total culture) are presented here. Cells were immunolabelled with Alexa Fluor 488 phalloidin for F-actin in green. Hoechst 33342 was used to stain the nuclei in blue. Images were taken at the same X-Y location but on different Z-planes. F-actin is expressed in both Calu-3 cells (A) and HBSMCs (B). Scale bar = 200µm.

A B

A B

Figure 5.20 Polar graphs showing frequency distribution of the angle of HBSMC alignment with respect to the inlet-outlet axis of the microfluidic channel. HBSMCs were co-cultured with Calu-3 cells on opposing sides of an ECM

hydrogel suspended inside a microfluidic device for 7 days. Radial (horizontal) axis represent the frequency of angle of alignment. Each bars represent a 100 range. Arrows point to the outlet of the microfluidic channel. Polar graphs represent frequency distribution for one representative sample on Day 4 (A) and Day 7 (B) of co-culture. A narrower distribution is observed for 7-day co-culture compared to a 4-day co-culture (n = 134). Mean alignment angles and standard deviations of each system are indicated on the top right corner of each graph for each system.

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 78° SD = 43°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73° SD = 21°

Mean = 73°



seeded on Day 0 and HBSMCs were seeded on Day 14. Representative images of co-culture on Day 4 (18-day total culture) and 7 (21-day total culture) are presented here. Cells were immunolabelled with mouse anti-α- smooth muscle actin mAb for α-smooth muscle actin (α-SMA) in red (A, B) and Rabbit anti-ZO-1 pAb for tight junction proteins in green (C, D). Hoechst 33342 was used to stain the nuclei in blue. Image pairs A&C and B&D were taken at the same X-Y location but on different Z-planes. α-SMA were expressed in HBSMCs and positive expression on ZO-1 proteins were observed in Calu-3 cells for both Day 4 and 7. Co-staining for α- SMA and ZO-1 showed no double positive cells. Scale bar = 200µm.

A B

C D


5.3 Discussion

Microfluidics offers a platform for in vitro model development that is robust enough to

incorporate complex tissue structures of the in vivo microenvironment and allow integration of

on-chip functionalities to simulate physiologic conditions, in its design. Increasing evidence

seems to suggest that these factors have a great influence on tissue engineering and tissue

development. In addition, using arrayable platforms allow us to increase throughput studies

while significantly reducing reagent consumption and device footprint. We demonstrate here the

use of our proposed arrayable PMMA microfluidic device to conduct several preliminary studies

to better understand the behavior of lung airway cells, Calu-3 cells and HBSMCs, in our system.

5.3.1 Calu-3 Cell Adhesion

At first, we showed adhesion preferences of Calu-3 cells on four different solutions of ECM

hydrogel inside a PMMA microfluidic device. The goal was to find an optimal solution that would

promote adhesion and proliferation of these cells into confluent monolayers. We used type 1

collagen as part of our study, which is know to be the most abundant fibrous protein within the

interstitial ECM that plays many roles including providing tensile strength, regulating cell

adhesion and supporting direct tissue development (Frantz, Stewart, and Weaver 2010). We

also used Matrigel, a commercially available preparation of basement membrane proteins, as

part of our study. Laminin, a major constituent of Matrigel, reportedly plays an essential role as a

biochemical and biomechanical inducer in polarization and tissue-like morphogenesis of

epithelial cells (Yu et al. 2008).

Although numerous studies in the past have shown that Calu-3 cells adhere, proliferate and

form confluent monolayers well on Col-1 coated substrates (Foster et al. 2000a) (Florea et al.

2003), (Zhang et al. 2016), not many studies have been conducted to show the effects of

Matrigel on Calu-3 epithelial cell adhesion.


In our study, we showed the influence of more types of protein or protein mixtures on Calu-3 cell

culture. More cells appeared to remain attached on hydrogel solutions with Col-1 or Col-1 in its

compositions on day 2. However, by Day 7 cells appeared to have detached from the Col-1

hydrogel and the hydrogel mixture with high concentrations of Col-1. Quantitative data showed a

significant decrease in area covered by adhered Calu-3 cells between Day 2 and 7 for these two

hydrogel solutions. This is in contrast to previous studies that have shown good Calu-3 cell

adhesion behavior to Col-1 coated substrates. Since cells actively sense and respond to both

chemical and physical stimuli from the environment, it is possible that for our system, aside from

integrin signalling between cell and ECM that are typically thought to influence cell adhesion,

other factors such as hydrogel stiffness, surface topography and growth factors in the

supplemented media are at play. A study conducted by Sivathanu et al., investigated the effect

of type 1 collagen gel thickness on Calu-3 cell growth, in which they discovered that thinner gels

(10 – 40 µm) promoted faster growth into confluent monolayers because the cells can sense the

stiff substrate beneath the gel more easily when gels are thinner (Sivathanu and Kamm 2013).

In keeping with this discovery, since Calu-3 cells in our system were seeded on hydrogels that

were approximately 500 µm thick, it is possible that cells in our system did not adhere well to

Col-1 or the composite hydrogel with a high concentration of Col-1 due to a lower stiffness of the

hydrogel than the cells prefer. Further investigation is required to better understand the impact

of Col-1 stiffness in Calu-3 epithelial adhesion and may be the subject of later studies. However,

for now it is not enough evidence to completely rule out Col-1 from this study because the

highest confluence of Calu-3 cells was found on a hydrogel mixture with Col-1 as one of its

constituents but with a high concentration of Matrigel. This results seems to suggest that

Matrigel has an effect on Calu-3 cell adhesion. But it is unexpected since we noticed that on day

2, adhesion behavior of cells on Matrigel was poor compared to the other hydrogels or mixtures

tested and did not change significantly between day 2 and 7, as confirmed by quantitative data.

This suggests that perhaps Matrigel alone is not sufficient to promote cell adhesion for a


long-term culture and that Col-1 has a stronger influence on cell adhesion when combined with

a high concentration of Matrigel. It is possible that the addition of Matrigel makes for a stiffer

hydrogel composite when polymerized, thus promoting Calu-3 cell adhesion. However, further

investigation is required to understand the influence of Matrigel or its protein constituents on

Calu-3 cell adhesion.

5.3.2 HBSMC Cell Adhesion

Next we examined adhesion preferences of HBSMCs to the same ECM hydrogels tested for

Calu-3 epithelial cells. In the past, cell-culture based studies of ASM have shown interstitial

ECM proteins such as Col-1 promote proliferation and the expression of less contractile

phenotype while basement membrane proteins stimulate expression of a differentiated

contractile phenotype and inhibits proliferation (Yamamoto, Yamamoto, and Noumura 1993), (Li

et al. 1994). This is thought to be due to the effect of laminin from Matrigel that has been

previously shown to inhibit airway smooth muscle cell growth (Hirst, Twort, and Lee 2000). In

line with these findings, we found that HBSMCs seeded on Matrigel did not adhere and spread

over a 7-day culture period compared to other hydrogels tested. This combined with the results

obtained from Calu-3 adhesion study, which showed poor adhesion to Matrigel prompted the

decision to eliminate Matrigel alone from further consideration in this study. The remaining

results showed that HBSMCs adhered and spread on Col-1 and the other hydrogel composites.

No significant differences in adhesion behavior, confirmed by quantitative data of adhered cells

per mm2 of substrate, between the three remaining hydrogel types were observed for both day-2

and day-7 data. In addition, cells appeared to grow and spread well over the 7-day culture

period for all three hydrogels examined. This is interesting, considering that the hydrogel

composite with Col-1 and a high concentration of Matrigel, ie. high laminin content, should

inhibit cell growth compared to the Col-1 hydrogel or the composite with high concentration of

Col-1 and low concentration of Matrigel, according to previous studies. A possible explanation


for this observation is that the primary integrin receptors, more specifically α1 β1 and α2 β2, that

mediate growth inhibition effects of laminin also interact with Col-1 (Thyberg 1996). Thus these

interactions may be counteracting the effects brought on by Laminin on the growth of HBSMCs.

From the results gathered so far we have decided to choose hydrogel composite [Col-1(3µg/µL)

& Matrigel (6µg/µL)], based on the highest adhesion yield from both cell types. Results from this

cell adhesion study suggests that both Col-1 and the protein constituents of Matrigel have roles

in promoting cell adhesion and growth of both Calu-3 cells HBSMCs. It is unclear from the

evidence gathered so far whether the impact of these proteins on these cells are physically or

chemically exclusive or a combination of both. It is important to answer these questions because

the exact composition of proteins in Matrigel is often unknown and as a result the cell adhesion

behavior may vary from batch to batch. Thus, future studies on the impact of gel stiffness and

the individual proteins in Matrigel on the adhesion behavior of each cell type may need to be

conducted to gain deeper understanding and improve reproducibility of outcomes using this

system.

5.3.3 Cell Seeding Density Study

The purpose of the next study was to determine the culture parameters such as cell seeding

density and culture time to obtain a confluent monolayer of Calu-3 epithelium and to reach full

confluence of aligned HBSMCs when cultured on a suspended hydrogel. In this study different

Calu-3 cell seeding densities led to noticeably different seeding efficiencies. For the proposed

device design, Calu-3 cells cultured at seeding densities 4000 cells/µL (~ 7800 cells/mm2) and

above formed cell clusters yielding a lower seeding efficiency. In addition, culture time for Calu-3

cells on our previously selected suspended ECM hydrogel extended beyond 2 weeks compared

to a culture on the same hydrogel that was supported by a flat PMMA substrate. A possible

explanation for this may be that cells can sense the stiff PMMA surface underneath the hydrogel


for the “non-suspended” case and as a result adheres and grows to confluence faster compared

to the “suspended hydrogel” case where large regions of the hydrogel are unsupported. Protein

losses in these unsupported regions due to channel washes during media replacement may

occur leading to a loss of hydrogel structural integrity and add to the effect of lower stiffness the

cell may experience from the hydrogel itself. Although in previous chapters the device design

has been optimized to provide sufficient support for the suspended hydrogel while maximizing

the exposed (free) surface of the hydrogel for cell-ECM contact, this does not completely

eliminate the low stiffness effect of these hydrogels.

Nevertheless, a confluent monolayer with area coverage above 80% was formed for seeding

density 3000 cells/µL (~ 5900 cell/mm2) by the second week of culture. Further studies may be

conducted to increase adhesion efficiency in the future but for now we have chosen two weeks

as the culture time for Calu-3 cells to grown into confluent monolayer, when seeded at 3000

cells/µL based on the results gathered here. One approach to investigate further into seeding

efficiencies may be to try different seeding patterns such as multiple seeding instances to

collectively allow more time for cells to attach. Other factors that may influence adhesion

efficiency are hydrogel stiffness and channel design, the impact of which may also be studied.

5.3.4 Monoculture and Co-culture Characterization

Our analysis of barrier properties of Calu-3 monolayers revealed the presence of TJ proteins

ZO-1 on and beyond day-14 of culture in both monoculture and co-culture systems. It is an

important preliminary finding in moving towards validating tissue functionality of this in vitro

model because TJ’s make vital contributions to the barrier properties of the bronchial epithelium.

Epithelium damages, opening of TJs and increased epithelial barrier permeability are common

in inflammatory conditions of the airways which lead to lung diseases (Foster et al., 2000).

Although further permeability studies may need to be conducted to test barrier function, the


presence of TJ proteins are positive indicators that TJs are present in these reconstituted

epithelium, and demonstrates potential for this in vitro model to be used in barrier integrity

studies of the airway epithelium.

HBSMCs in both monocultures and co-cultures expressed F-actin and α-SMA, which is also an

important preliminary finding in the context of in vitro model development because of their

crucial roles as a marker of various structural functions like contractility and motility. As

mentioned before airway remodelling as a result of structural changes of smooth muscle tissue

are linked with pathogenesis of lung diseases like asthma. The exact mechanisms of airway

remodelling in these diseases are still the subject of many studies and thus the expression of

these proteins in our system indicate there is potential for these types of studies to be conducted

in our system.

Alignment studies revealed that with culture time and higher cell densities, HBSMCs tended to

preferentially align along the length of the channel. An explanation for this observation may be

that the cells prefer to align along the path of least resistance which in this case was along the

length of the channel. If this is the case, then we may be able to control the alignment of these

cells with channel designs. However, this explanation is speculative and more experiments will

need to be conducted to study the effects of channel design on HBSMC alignment. To recreate

the similar morphology of aligned bands smooth muscle tissues that wrap around the

bronchioles and bronchi of the human lung, we noticed that longer culture period and higher cell

densities yields a narrow distribution of alignment angles (ie. more aligned cells).

The goal of this chapter was to finalize of cell culture parameters required to develop a

co-culture system with a Calu-3 epithelium and confluent culture of HBSMCs on opposing sides

of a suspended ECM. Furthermore, the goal was to perform preliminary assessment of the

cultures grown in our devices. We have shown that using our proposed PMMA microfluidic

device, we were able to create an array of monoculture and co-culture systems in a single


device and use the device to conduct several studies to better understand the behavior of

Calu-3 cells and HBSMCs in our system. The results of theses studies, summarized in this

chapter, revealed critical information on what culture parameters are required for the formation

of a monolayer epithelium using Calu-3 cells and a confluent layer of smooth muscle cells using

HBSMCs on opposing sides of a suspended ECM hydrogel. We have shown that not only do

these cells survive long term static cultures in our PMMA microfluidic device but they also

demonstrate some important characteristics of a functional bronchial epithelium. Although not

much work has been done to characterize the co-culture systems in this study and additional

examination is required to demonstrate further functionality of this device, the preliminary

assessment results show great potential of this device as an in vitro lung airway model.

97

6.0 Conclusions and Recommendations

6.1 Summary

A PMMA microfluidic device for co-culturing airway epithelial cells (Calu-3) and HBSMCs has

been designed, fabricated and employed in a number of preliminary investigations to assess its

fitness as an airway in vitro model. The primary aspect of the design involves a suspended ECM

hydrogel that is more physiologically relevant than current ECM mimics used in microfluidic

airway modelling and that can support long-term culture of the two airway cell types. The design

is set up to potentially accommodate ALI culture inside the microfluidic device. We have chosen

a microfabrication technique that allows ultra-rapid prototyping and a thermoplastic-based

platform that is suitable for making arrays of culture systems in a single microfluidic device. This

not only accelerated our progress in running through numerous design iterations but also

enabled us to gather a wealth of information in a single experiment at a time. To prepare our

device for cell culture, we first looked into design limitations associated with suspending a

hydrogel inside our proposed device. Principles of suspended microfluidics were investigated

and extended to produce high (width to height) aspect ratio fluid suspensions. An analytical

model was proposed to predict the limitations of channel design parameters based on fluidic

properties.

6.0 Conclusions and Recommendations 98

As part of preliminary investigations, the device was used to investigate an optimal hydrogel

composition that promotes cell attachment and proliferation. Col-1, Matrigel and their mixtures

were used for this investigation and revealed that a hydrogel mixture with high concentration of

Matrigel and low concentration of Col-1 yielded the highest cell adhesion output. Further, the

device and the selected hydrogel mix was used to determine critical cell culture parameters

required to form a confluent Calu-3 monolayer epithelium and a confluent band of aligned

HBSMCs inside the microfluidic device. We found that for the proposed device design, Calu-3

cells cultured on a suspended ECM hydrogel took between 2 to 3 weeks to form a confluent

monolayer covering above 85% of the ECM hydrogel surface. We also found that HBSMCs

were less sensitive to culture conditions like cell seeding density and culture time than Calu-3

cells. HBSMCs can be grown to confluence very easily by controlling seeding density or culture

time.

Finally, Calu-3 monolayers were evaluated for the expression of tight junction proteins that are

involved in forming tight cell-to-cell junctions which then control the movement of molecules and

ions across the epithelium. ZO-1 TJ proteins were expressed on Day 14 and 21 of culture, even

under co-culture conditions. HBSMCs were evaluated for their expression of F-actin and α-SMA,

both of which have roles in important cellular process such as motility and contractile functions.

Expression of both proteins were found in both monoculture and co-culture systems.

Furthermore, in both monoculture and co-culture systems highly aligned HBSMCs were found

with increasing culture time.

In conclusion these findings demonstrate the usefulness of the proposed platform not only for its

ability to produce results in a high throughput fashion but also in its ability to accommodate

long-term cell culture, maintenance and characterization protocols without changing the

characteristic phenotypes of the two cell types cultured in this device. More importantly,

preliminary results show that there is great potential in using this platform to develop a


well-functioning, more physiologically relevant airway in vitro model for conducting disease

mechanism and drug development studies.

6.2 Recommendations

Given our current findings, a few studies may be conducted immediately to expand the

usefulness of our system.

In order to standardize the way device design parameters are obtained to suspend any

hydrogels or liquids with various fluidic properties:

1. An investigation into developing a more realistic numerical model for pressure-driven

suspended microflow may be conducted

To enhance our understanding of the specific biochemical and biophysical factors of ECM

properties that influence cell adhesion behavior of the two cells used in developing this model:

1. A more thorough study on the specific effects of each ECM proteins and their

concentration on cell adhesion behavior may be conducted

2. Also an investigation into the effects of substrate (hydrogel) stiffness on cell adhesion

behavior in our device may be conducted.

To shorten culture time of Calu-3 cells in our proposed microfluidic device:

1. The effects of cell seeding patterns on monolayer formation of Calu-3 cell may be

examined

2. The effects of device design on monolayer formation of Calu-3 cells may also need to be

examined

To further characterize the barrier properties of the reconstituted epithelium:


1. A permeability study may be conducted to investigate the barrier functions of the Calu-3

epithelium

The next steps in providing a more representative model of the airway epithelium would be to

culture Calu-3 cells at an ALI:

1. In order to culture Calu-3 cells at an ALI, culture parameters such as from media

supplements required to sustain this type of culture, and the time timing of culture steps

will need to be examined.

101

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107

Appendix A

A1.0 Theoretical Approach to Characterizing Suspended Microflow

with Applied Pressure

The type of systems under consideration involve pressure-driven microflows within open

microstructures consisting of multiple air or immiscible fluidic interfaces

A1.1 Free Energy Calculation

To predict the onset of flow we need to evaluate the free energy of the system over time

Since our system is a constant pressure and temperature system we may utilize the Gibbs

free energy equation to calculate the free energy of our system.

Gibbs free energy equation predicts the spontaneity of a process, in other words, it

determines whether or not a process (in this case fluid flow) can occur

𝑑𝐺 = 𝛾𝑑𝐴 − 𝑝𝑑𝑉 − 𝑠𝑑𝑇

where

𝛾 = surface tension

A = liquid surface area

V = liquid volume

T = temperature

s = entropy

p = liquid pressure

dG = change in Gibbs free energy (measuring surface energy)

The equation above determines the change in Gibbs free energy of a system at constant

temperature and pressure. A negative change in Gibbs free energy indicates the process is

spontaneous and may proceed forward, whereas a positive change means the process is

non-spontaneous and may proceed in the reverse direction over time. A zero change is

representative of a system at equilibrium.

Since in our case we are only concerned with whether or not fluid flow may occur, we can

modify the above equation to search for conditions where dG <0,

Appendix A 108

Assuming negligible temperature change and realizing that the volume of liquid is

constantly increasing, Gibbs free energy equation simplifies to

𝒅𝑮 = 𝜸𝒅𝑨 − 𝑷𝒅𝑽 < 0

The system under consideration may consists of multiple air or immiscible fluidic

interfaces. Therefore, the above equation is more accurately represented as

∑ 𝜸𝒊

𝒊

𝒅𝑨𝒊

𝒅𝑽− 𝑷 < 𝟎

where

∑ 𝜸𝒊

𝒊

𝒅𝑨𝒊

𝒅𝑽= 𝜸𝑺𝑮

𝒅𝑨𝑺𝑮

𝒅𝑽+ 𝜸𝑺𝑳

𝒅𝑨𝑺𝑳

𝒅𝑽 + 𝜸𝑳𝑮

𝒅𝑨𝑳𝑮

𝒅𝑽

SG = solid gas ; SL = solid liquid; LG = liquid gas

As the system evolves towards equilibrium, the solid-gas interface is replaced by

the solid-liquid interface 𝒅𝑨𝑺𝑳

𝒅𝑽= −

𝒅𝑨𝑺𝑮

𝒅𝑽

∑ 𝜸𝒊

𝒊

𝒅𝑨𝒊

𝒅𝑽= −𝜸𝑺𝑮

𝒅𝑨𝑺𝑳

𝒅𝑽+ 𝜸𝑺𝑳

𝒅𝑨𝑺𝑳

𝒅𝑽 + 𝜸𝑳𝑮

𝒅𝑨𝑳𝑮

𝒅𝑽

To simply the above equation, we consider Young's Equation which provides a

relationship between the solid-liquid interfacial tension , solid surface free energy,

contact angle and the liquid surface tension

𝜸𝑳𝑮 𝐜𝐨𝐬 𝜽 = 𝜸𝑺𝑮 − 𝜸𝑺𝑳

Equation 1 then becomes

𝜸𝑳𝑮 ∑ [− 𝐜𝐨𝐬 𝜽𝒊

𝒅𝑨𝑺𝑳,𝒊

𝒅𝑽]

𝒊

+ 𝜸𝑳𝑮

𝒅𝑨𝑳𝑮

𝒅𝑽 − 𝑷 < 0

It should be noted that the above equation states that a larger solid-liquid interface

will result in energy reduction and larger ALI with result in energy increase

To calculate 𝒅𝑨𝑺𝑳,𝒊

𝒅𝑽 and

𝒅𝑨𝑳𝑮

𝒅𝑽 we focus on the contours (perimeter) of the

interface (solid-liquid and liquid-gas) under condition. Pw and Pf are the wetted and

free perimeter respectively.

Appendix A 109

𝒅𝑨𝑺𝑳,𝒊 = 𝒅𝒙 ∗ 𝒑𝒘

𝒅𝑨𝑳𝑮,𝒊 = 𝒅𝒙 ∗ 𝒑𝒇

Therefore, the equation below calculates the free energy of a suspended microflow

system with applied pressure

𝜸𝑳𝑮 ∑ [− 𝐜𝐨𝐬 𝜽𝒊

𝒅𝒙 ∗ 𝒑𝒘,𝒊

𝒅𝑽]

𝒊

+ 𝜸𝑳𝑮

𝒅𝒙 ∗ 𝒑𝒇

𝒅𝑽 − 𝑷 < 0

A1.2 Analytical Model of Pressure Driven Suspended Microflow

𝒑𝒇 − 𝑷

𝜸𝑳𝑮

𝒅𝑽

𝒅𝒙 < ∑[ 𝐜𝐨𝐬 𝜽𝒊 𝒑𝒘𝒊]

𝒊

The above equation represents an analytical model for a pressure driven

suspended microflow in a composite microfluidic channel. The model incorporates

design parameters, such as properties of the solid material, applied pressure and

channel geometry, that can be used to predict the onset of suspended microflow

For a system having homogeneous solid walls

𝒑𝒇

𝒑𝒘−

𝑷

𝒑𝒘 𝜸𝑳𝑮

𝒅𝑽

𝒅𝒙 < 𝐜𝐨𝐬 𝜽𝑺𝒐𝒍𝒊𝒅

The following equation can be used to compare surface energies of different solid

material and surface morphology

𝒑𝒇

𝒑𝒘−

𝑷

𝒑𝒘 𝜸𝑳𝑮

𝒅𝑽

𝒅𝒙− 𝐜𝐨𝐬 𝜽𝒔𝒐𝒍𝒊𝒅 < 0

Minimum pressure required to facilitate suspended microflow can also be determined

for cases where 𝑷𝒇

𝑷𝒘≥ 𝐜𝐨𝐬 𝜽𝒊

[𝒑𝒇

𝒑𝒘− 𝐜𝐨𝐬 𝜽𝒊] 𝒑𝒘 𝜸𝑳𝑮

𝒅𝒙

𝒅𝑽< 𝑷

𝒑𝒇 < ∑[ 𝐜𝐨𝐬 𝜽𝒊 𝒑𝒘𝒊]

𝒊

w

w

w

w

w

w

w

w

w

w

w

Appendix A 110

A2.0 Dynamic Approach of Pressure Driven Suspended Microflow

between Parallel Plates

Here we present an analytical approach for describing the dynamics of pressure driven, suspended flow between parallel plates

A2.1 Description of the Physical Problem

Fluid flow between the parallel plates is induced by surface tension and applied pressure at the inlet of the channel

Forces acting on the fluid body under consideration include viscous drag (resisting the motion of fluid), pressure force and capillary forces (both of which assisting fluid motion). There is negligible external body forces (eg. gravity since dimensions are below capillary length)

Flow is newtonian and laminar. Assuming that a uniform cross-section (rectangular) is maintained along the length of channel.

Figure A1.1 Schematic of pressure driven suspended microflow between parallel

plates.

Flow

Flow

Flow

Flow

Flow

Flow

Flow

w

w

w

w

w

w

w

h

h

h

h

h

h

h

Appendix A 111

A2.2 The General Governing Equation of Fluid Motion

Let's assume that the instantaneous axial length occupied by fluid is x and the

governing equation of fluid motion derived from force balance is as follows

𝒎𝒅( 𝒗𝒂𝒗𝒈)

𝒅𝒕 = 𝑭𝒑𝒓𝒆𝒔𝒔𝒖𝒓𝒆 + 𝑭𝒄𝒂𝒑 − 𝑭𝒅𝒓𝒂𝒈

The LHS represents the transient inertial term and the RHS includes pressure,

capillary and drag force

Expanding the LHS and assuming that a constant shape is advancing,

𝒎𝒅( 𝒗𝒂𝒗𝒈)

𝒅𝒕= 𝝆[(𝒉 𝒘) 𝒙(𝒕)]

𝒅𝒗𝒂𝒗𝒈

𝒅𝒕

A2.3 Pressure, Capillary and Drag Force Calculation

See section 3 for the profile of a pressure driven suspended microflow between

parallel plates.

The pressure force of the system can be calculated as follows.

𝑭𝒑𝒓𝒆𝒔𝒔𝒖𝒓𝒆 = ∫𝝏𝒑

𝝏𝒙𝒅𝑽

𝑽 ∫ ∫ ∫

∆𝒑

𝑳 𝒅𝒙𝒅𝒚𝒅𝒛

𝑳

𝒉

𝟐

−𝒉

𝟐

𝒘

𝟐

−𝒘

𝟐

= ∆𝒑 𝒉 𝒘

The capillary force assuming the advancing contact angle of the fluid front equals

Young's contact angle

𝑭𝒄𝒂𝒑 = 𝒑𝒘𝜸 𝐜𝐨𝐬 𝜽 − 𝜸 𝒑𝒇 = 𝟐𝜸 [𝒉 𝐜𝐨𝐬 𝜽 − 𝒘 ]

The drag force assuming a poiseuille flow profile between the vertical walls

𝑭𝒅𝒓𝒂𝒈 = 𝟔𝝁𝒗𝒂𝒗𝒈

𝒘𝒙(𝒕)(𝒑𝒘) = 𝟏𝟐𝝁

𝒉

𝒘𝒙(𝒕)𝒗𝒂𝒗𝒈(𝒕)

A2.4 Force Balance

Substituting equation 2 and 5 into 1 we get

Appendix A 112

𝝆[(𝒉 𝒘) 𝒙(𝒕)]𝒅𝒗𝒂𝒗𝒈

𝒅𝒕= ∆𝒑 𝒉 𝒘 + 𝟐 𝜸 [𝒉 𝒄𝒐𝒔 𝜽 − 𝒘 ] − 𝟏𝟐𝝁

𝒉

𝒘𝒙(𝒕)𝒗𝒂𝒗𝒈(𝒕)

Rearranging

𝒅𝒗𝒂𝒗𝒈

𝒅𝒕−

∆𝒑 𝒉 𝒘

[𝝆(𝒉 𝒘) 𝒙(𝒕)]−

𝟐𝜸 [𝒉 𝐜𝐨𝐬 𝜽 − 𝒘 ]

𝝆[(𝒉 𝒘)𝒙(𝒕)]+

𝟏𝟐𝝁𝒉

𝝆[(𝒉 𝒘)𝒘] 𝒗𝒂𝒗𝒈(𝒕) = 𝟎

Substituting 𝒗𝒂𝒗𝒈(𝒕) = 𝒅𝒙

𝒅𝒕 we get

𝒅𝟐𝒙

𝒅𝒕𝟐+ (

𝟏𝟐 𝝁

𝝆 𝒘𝟐)

𝒅𝒙

𝒅𝒕−

∆𝒑

𝝆 𝒙(𝒕) −

𝟐 𝜸 [ 𝐜𝐨𝐬 𝜽 − 𝒘𝒉

]

𝝆 𝒘

𝟏

𝒙(𝒕) = 𝟎

A2.5 Force Balance Neglecting the Inertial force

Neglecting the inertial term due to low Reynolds number (low flowrate), the force

balance

∆𝒑 𝒉 𝒘 + 𝟐 𝜸 [𝒉 𝒄𝒐𝒔 𝜽 − 𝒘 ] − 𝟏𝟐𝝁𝒉

𝒘𝒙(𝒕)𝒗𝒂𝒗𝒈(𝒕) = 𝟎

A2. 6 Velocity Profile

Velocity relationship with respect to the axial length (direction of flow)

𝑽 = 𝒅𝒙

𝒅𝒕=

∆𝒑𝒘𝟐

𝟏𝟐𝝁𝒙(𝒕)+

𝜸𝒘[𝒄𝒐𝒔𝜽 − 𝒘/𝒉]

𝟔𝝁𝒙(𝒕)

To obtain a time dependent velocity profile we integrate equation 8 with respect

to time which yields an expression for distance traveled with respect to time.

𝒙(𝒕)𝒅𝒙 = [∆𝒑𝒘𝟐

𝟏𝟐𝝁+

𝜸𝒘[𝒄𝒐𝒔𝜽 − 𝒘/𝒉]

𝟔𝝁] 𝒅𝒕

𝒙(𝒕) = √[∆𝒑𝒘𝟐

𝟔𝝁+

𝜸𝒘

𝟑𝝁[𝒄𝒐𝒔𝜽 −

𝒘

𝒉]] 𝒕

The time dependent velocity profile of the flow is deduced by finding the derivative

of equation 9.

Appendix A 113

𝑽 = √∆𝒑𝒘 𝟐

𝟐𝟒𝝁+

𝜸𝒘

𝟏𝟐𝝁[𝒄𝒐𝒔𝜽 −

𝒘

𝒉] √

𝟏

𝒕

A3.0 Flowrate to Pressure drop

A3.1 Relationship Between Pressure Drop and Flowrate in a

Microfluidic Channel

Pressure driven, steady state flow through a constant cross-sectional shape can be characterized by Poiseuille flow. Constant pressure drop results in a constant flow rate, Q.

∆𝒑 = 𝑹𝒉𝒚𝒅 ∗ 𝑸

where ∆𝑝 , is the pressure drop along the channel, Q is the flow rate, 𝑅ℎ𝑦𝑑 is the

proportionality factor called hydraulic resistance; central concept in characterizing

and designing microfluidic channels in lab-on-a chip systems.

𝑹𝒉𝒚𝒅 = ∆𝒑

𝑸

Volumetric flow rate relates to the velocity profile equation in the following way

𝑸 = ∫ 𝒅𝒚 𝒅𝒛 𝒗𝒙(𝒚, 𝒛) = 𝑨𝒄𝒗𝒂𝒗𝒈

𝒄

where Ac is the cross-sectional area of the fluid flow and 𝑣(𝑥, 𝑦) is the velocity profile of the fully developed flow and vavg is the average velocity denoted by

𝒗𝒂𝒗𝒈 = (𝟏

𝑨𝒄) ∫ 𝒅𝒚 𝒅𝒛 𝒗𝒙(𝒚, 𝒛)

𝒄

A3.2 Volumetric Flowrate Calculation from Velocity Profile

𝑸 = ∫ 𝒅𝒚 𝒅𝒛 𝒗𝒙(𝒚, 𝒛)

𝒄

Appendix A 114

𝑸 = ∫ ∆𝒑𝒘𝟐

𝟖𝑳𝝁{[𝟏 − 𝟒

𝒚𝟐

𝒘𝟐

] + ∑ [𝟑𝟐 (−𝟏)𝒏

𝝀𝒏𝟑

𝐜𝐨𝐬 (𝝀𝒏𝒚𝒘) 𝐜𝐨𝐬𝐡 (𝝀𝒏

𝒛𝒘)

𝒄𝒐𝒔𝒉 (𝝀𝒏𝒉𝟐𝒘 )

]

∞

𝒏=𝟎

} 𝒅𝒚 𝒅𝒛

𝒄

∫ ∆𝑝𝑤2

8𝐿𝜇{[1 − 4

𝑦2

𝑤2

]} 𝑑𝑦 𝑑𝑧 = ∆𝑝𝑤3

8𝐿𝜇{ℎ − (

ℎ3

3𝑤2)}

𝒄

∆𝑝𝑤2

8𝐿𝜇∫ {∑ [

𝟑𝟐 (−𝟏)𝒏

𝝀𝒏𝟑

𝐜𝐨𝐬 (𝝀𝒏𝒚𝒘

) 𝐜𝐨𝐬𝐡 (𝝀𝒏𝒛𝒘

)

𝒄𝒐𝒔𝒉 (𝝀𝒏𝒉𝟐𝒘 )

]

∞

𝒏=𝟎

} 𝑑𝑦 𝑑𝑧

𝑐

=16∆𝑝𝑤4

𝐿𝜇∑ [

(−1)𝑛

((2𝑛 + 1)𝜋)5 sin (

(2𝑛 + 1)𝜋

2) tanh (

(2𝑛 + 1)𝜋ℎ

2𝑤)]

∞

𝑛=0

𝑄 = ∆𝑝𝑤3

8𝐿𝜇{ℎ − (

ℎ3

3𝑤2)}

+16∆𝑝𝑤4

𝐿𝜇∑ [

(−1)𝑛

((2𝑛 + 1)𝜋)5 sin (

(2𝑛 + 1)𝜋

2) tanh (

(2𝑛 + 1)𝜋ℎ

2𝑤)]

∞

𝑛=0

𝑄 = ∆𝑝ℎ𝑤3

8𝐿𝜇{1 −

ℎ2

3𝑤2

+ (128𝑤

ℎ) ∑ [

(−1)𝑛

((2𝑛 + 1)𝜋)5 sin (

(2𝑛 + 1)𝜋

2) tanh (

(2𝑛 + 1)𝜋ℎ

2𝑤)]

∞

𝑛=0

}

A3.3 Flowrate Simplification

Let

𝛼 =ℎ𝑤3

8𝜇{1 −

ℎ2

3𝑤2+ (

128𝑤

ℎ) ∑ [

(−1)𝑛

((2𝑛 + 1)𝜋)5

sin (

(2𝑛 + 1)𝜋

2) tanh (

(2𝑛 + 1)𝜋ℎ

2𝑤)]

∞

𝑛=0

}

Evaluate summation of series by checking for convergence of

Appendix A 115

∑ [ (−1)𝑛

((2𝑛 + 1)𝜋)5 sin (

(2𝑛 + 1)𝜋

2) tanh (

(2𝑛 + 1)𝜋ℎ

2𝑤)]

∞

𝑛=0

Take out constant

1

𝜋5∑ [

(−1)𝑛 sin ((2𝑛 + 1)𝜋

2 ) tanh ((2𝑛 + 1)𝜋ℎ

2𝑤 )

(2𝑛 + 1)5 ]

∞

𝑛=0

Apply Squeeze Theorem

−1 ≤ sin ((2𝑛 + 1)𝜋

2) ≤ 1

∑ [ (−1)𝑛 (−1) tanh (

(2𝑛 + 1)𝜋ℎ2𝑤 )

(2𝑛 + 1)5 ]

∞

𝑛=1

≤ ∑ [ (−1)𝑛 sin (

(2𝑛 + 1)𝜋2 ) tanh (

(2𝑛 + 1)𝜋ℎ2𝑤 )

(2𝑛 + 1)5 ]

∞

𝑛=0

≤ ∑ [ (−1)𝑛 (1) tanh (

(2𝑛 + 1)𝜋ℎ2𝑤 )

(2𝑛 + 1)5 ]

∞

𝑛=0

Check convergence of

∑ [ (−1)𝑛 (1) tanh (

(2𝑛 + 1)𝜋ℎ2𝑤 )

(2𝑛 + 1)5 ]

∞

𝑛=0

Alternating Series Test

if lim𝑛 →∞

𝑏𝑛 = 0

then ∑(−1)𝑛 𝑏𝑛 𝑎𝑛𝑑 ∑(−1)𝑛−1 𝑏𝑛 𝑐𝑜𝑛𝑣𝑒𝑟𝑔𝑒𝑠

𝑏𝑛 = (1) tanh (

(2𝑛 + 1)𝜋ℎ2𝑤 )

(2𝑛 + 1)5

Appendix A 116

lim𝑛 →∞

tanh ((2𝑛 + 1)𝜋ℎ

2𝑤 )

(2𝑛 + 1)5 = 0

Therefore

∑ [ (−1)𝑛

((2𝑛+1)𝜋)5 sin (

(2𝑛+1)𝜋

2) tanh (

(2𝑛+1)𝜋ℎ

2𝑤)]∞

𝑛=0 =1

𝜋5[0] = 0

Alpha simplified

𝛼 =ℎ𝑤3

8𝜇{1 −

ℎ2

3𝑤2 + (128𝑤

ℎ𝜋5 ) ∗ 0} = ℎ𝑤3

8𝜇{1 −

ℎ2

3𝑤2}

𝑄 = ∆𝑝ℎ𝑤3

8𝐿𝜇{1 −

ℎ2

3𝑤2}

Therefore the hydraulic resistance and the pressure drop expression is

𝑅ℎ𝑦𝑑 = ∆𝑝

𝑄=

𝐿

ℎ𝑤3

8𝜇{1 −

ℎ2

3𝑤2}

∆𝑝 = 𝑄𝐿

𝛼=

𝑄𝐿

ℎ𝑤3

8𝜇 {1 −ℎ2

3𝑤2}

Appendix A 117

Day 4 System 1 System 2

System 3 System 4

Mean = 84° SD = 37°

Mean = 94° SD = 40°

Mean = 94° SD = 44°

Mean = 84° SD = 39°

A4.0 Frequency distribution of HBSMC alignment angle

A

A

A

A

A

A

A

A


System 3 System 4

Mean = 96° SD = 14°

Mean = 92° SD = 17°

Mean = 99° SD = 15°

Mean = 95° SD = 18°

B

Appendix A 118

Figure A4.1 Polar graphs showing frequency distribution of the angle of HBSMC alignment with respect to the inlet-outlet axis of the microfluidic channel. HBSMCs were culture on the underside of an ECM hydrogel suspended inside a microfluidic device and images were obtained on Day 4, 7 and 14 of culture. Radial (horizontal) axis represent the frequency of angle of alignment. Each bars represent a 100 range. Arrows point to the outlet of the microfluidic channel. Polar graphs represent frequency distribution for four systems assessed on Day 4 (A), Day (B) and Day 14 (C) of culture (n = 134). Mean alignment angles and standard deviations of each system are indicated on the top right corner of each graph for each system.


System 3 System 4

Mean = 72° SD = 11°

Mean = 96° SD = 12°

Mean = 96° SD = 12°

Mean = 88° SD = 13°

C

Appendix A 119

System 3 System 4

Day 4

System 1 System 2

Mean = 78° SD = 43°

Mean = 86° SD = 50°

Mean = 81° SD = 47°

Mean = 81° SD = 48°

A

Figure A4.2 Polar graphs showing frequency distribution of the angle of HBSMC alignment with respect to the inlet-outlet axis of the microfluidic channel. HBSMCs were co-cultured with Calu-3 cells on opposing sides of an ECM hydrogel suspended inside a microfluidic device and images were obtained on Day 4 and 7 of co-culture. Polar graphs represent frequency distribution for one representative sample on Day 4 (A) and Day 7 (B) of co-culture (n = 134). Mean alignment angles and standard deviations of each system are indicated on the top right corner of each graph for each system.

System 3 System 4

Day 7

System 1 System 2

Mean = 73° SD = 21°

Mean = 76° SD = 16°

Mean = 68° SD = 19°

Mean = 76° SD = 17°

B

Appendix A 120

A5.0 Device fabrication and Cell Culture Protocol

A5.1 Device Fabrication Protocol

Softwares

1. Solidworks

2. SprutCAM

Equipment

1. Tormach PCNC 770 vertical milling machine

2. Carver automatic hydraulic laboratory press

Materials

1. PMMA sheets

2. 1/16” (1.5875 mm) (Part # 01982156) endmill

3. 3/64″ (1.1906 mm) (Part # 07765431) endmill

4. 0.15″ (381 μm) (Part # 37289501) endmill

5. 0.020″ (508 μm) (Part # 37290525) endmill

6. Transparency film (215 mm x 279 mm)

7. Acetone

8. Double-sided tape

9. 70% Ethanol in distilled water

10. 99% Ethanol

11. Soap

12. Compressed air

13. Flat steel plates (153 mm X 153 mm)

14. Pipette (200µL)

15. Pipette tip (200 µL)

16. FisherbrandTM label tape

Procedure

Microfabrication

1. Design features into each layers of the device and assemble the layers in Solidworks.

Save file as a (. SLDASM) file and (. IGS) file for SprutCAM.

2. Prepare (.TAP) file using SprutCAM for CNC milling machine. Mill large channels and

ports with roughing waterline operation using the 1/16” endmill. Use roughing waterline

operation to mill hydrogel channels using 3/64” endmill and suspension channels using

0.015” endmill. Select 2D contouring operation and 3/64” endmill for carving out the

layers of the device. Use multiple passes for each operation to obtain smooth surface

finishing.

Appendix A 121

3. To mill device, clean the surface of the granite block attached to the milling machine

with acetone thoroughly.

4. Cut-out transparency film to a size slightly larger than the PMMA piece to be milled out.

5. Attach double-sided tape on one of the transparency film and place the taped side of

the film on the granite block. Ensure no air bubble are trapped between the tape and

the granite block.

6. Attach double-sided tape on one of the PMMA workpiece and place the workpiece on

the transparency film (sacrificial layer). Ensure no air bubble are trapped between the

tape and the film.

7. Load the (.TAP) file onto the milling machine computer and begin milling procedure.

Change endmill bits as required.

8. After milling, clean each layers of the device with laboratory soap and water to remove

coolant from the milling procedure, and dry off the milled pieces with compressed air.

9. Remove any residual double sided tape attached to the milled PMMA pieces using

fisherbrandTM label tape and rinse the PMMA pieces a second time with soap and

water.

10. Rinse the PMMA pieces with 70% ethanol in DI water and dry off using compressed air.

Bonding

1. Set carver automatic hydraulic laboratory press to 70oC and allow platens and the steel

plates to reach this temperature.

2. Load 100µL of 99% ethanol via pipetting in between two layers of PMMA and align the

pieces to mirror each other. Place the pieces in between steel plates.

3. Press the layers of PMMA using the automatic hydraulic laboratory at 1000 lbf for 1

min.

4. Remove bonded PMMA stacks let it cool down to room temperature. Repeat steps 2 to

4 for other layers of PMMA to be bonded.

5. After bonding, dry remaining solution inside the PMMA stacks with nitrogen gas and

leave stacks at room temperature overnight for all existing volatiles to evaporate.

A5.2 Cell Culture Protocol

Reagents

1. 70% ethanol in DI water 2. phosphate buffered saline (PBS) 3. 100 μg/mL bovine plasma fibronectin (FN) 4. Collagen type I ( Col-1) 5. Aliquoted Matrigel 6. Trypsin- EDTA (0.25%) 7. 4% paraformaldehyde (PFA) 8. Calcein AM 9. ethidium homodimer -1 10. Hoechst 33342 11. 0.1% Triton X-100 12. rabbit anti-ZO-1 pAb

Appendix A 122

13. mouse anti-α- SMA mAb 14. AlexaFluor 568 goat anti-rabbit IgG 15. ActinGreen 488 16. AlexaFluor 568 goat anti-mouse 17. Alexa Fluor 488 phalloidin . 18. 0.5N NaOH

Materials

1. 500 µL Eppendorf tube 2. Kimwipe 3. Eppendorf tube 4. 15 mL conical tube 5. Omnitray 6. single edge steel blade. 7. scalpel. 8. 1.2 mm thick 75 X 25 mm glass microscope slide 9. Hemacytometer

Equipment

1. Laboratory Incubator 2. Mini vortex mixer (Fisher Scientific) 3. Biological Safety Cabinet (BSC) 4. Aspirator 5. Laboratory Refrigerated Centrifuge (Fisher Scientific)

Procedure

Perform all tasks involving direct access to cell culture sample inside a BSC. Gel Loading

1. To prepare a 155 µL of hydrogel solution, mix 45 µL of (Col-I) and 9 µL of 0.5 N NaOH in

a 500 µL Eppendorf tube and set aside in a 4oC fridge for at least 1 hour. Thaw aliquoted

Matrigel in the 4oC fridge during this step as well.

2. Disinfect PMMA device by pipetting and aspirating 70% ethanol in DI water through the

compartments of the device two times.

3. Flush with 70% ethanol in DI water solution a third time and leave the solution in the

device for 20 min.

4. Aspirate 70% ethanol in DI water solution out and rinse with PBS three times.

5. Aspirate residual PBS solution from the device and leave the device inside the BSC for

at least 20min to evaporate remaining PBS volatiles.

6. Coat hydrogel channels with 100 μg/mL FN by dispensing 10 μL of FN solution into the

smaller channels in the middle layer of the device and incubate at room temperature for

at least 45 min.

7. Aspirate out FN from the device and contain the device in a sealed Omnitray.

Appendix A 123

8. Keep Omnitray in the -20 oC freezer for 15 min.

9. Mix 103 µL of thawed Matrigel into the Eppendorf tube containing Col-1 and NaOH

mixture prepared earlier and vortex for 10 to 15 sec. Keep Eppendorf tube icepack until

gel loading into the device.

10. After cooling the device move Omnitray to BSC and position the PMMA device for

hydrogel loading.

11. Tilt the device at a 45o angle towards the operator in an orientation such that the inlets

ports are above the outlet and dispense 10 µL of prepared ECM hydrogel solution into

the device through the inlet of the top compartment. Adjust dispensing speed according

to the rate at which the hydrogel solutions wet the walls of the channel via surface

tension.

12. After gel loading, place device back in the Omnitray. Place a sterile and saturated (with

sterile PBS) Kimwipe inside the Omnitray. Place Omnitray in a larger bioassay dish

containing 5 mL of sterile water and incubate at 37 °C, 5% CO2, for 2 hours for

polymerization.

13. After gel polymerization, dispense supplemented media of the respective cell type under

investigation in both the top and bottom compartments of each system in the device and

incubate at 37 °C, 5% CO2, for 3-6 hours.

Cell Seeding and Culture

1. Prepare cell suspension by removing old media out of Calu-3 cell or HBSMC flasks and

flushing with PBS.

2. Add Trypsin- EDTA (0.25%) at 37 °C to flasks for 10 min (Calu-3 cells) and for 2 min

(HBSMCs).

3. Dilute Trypsin- EDTA with respective supplemented media and suspended cells by

repeated pipetting of cell + media + Trypsin- EDTA (0.25%) solution up and down into

the flask.

4. Use this solution for cell counting using a hemacytometer.

5. Transfer of cell + media + Trypsin- EDTA (0.25%) solution to 15 mL conical tube and

centrifuge at (170 g; 7 min) for Calu-3 cells and at (220g ;3 min) for HBSMCs

6. Aspirate separated solution from conical tube and resuspend cells at 3000cells/μL for

Calu-3 cells and 500cells/μL for HBSMCs in fresh supplemented media. Suspende cells

by repeated pipetting of cell + media solution up and down into the conical tube.

7. Aspirate media from the top and bottom compartments of the device through the device

outlet ports.

8. Depending on the cell type under investigation, perform the following:

a. For Calu-3 cell seeding, dispense 80 μL of suspended cell solutions into top

compartment of the PMMA device through the inlet ports. Dispense 50 µL of fresh

supplemented media into the bottom compartment of the device through the inlet ports.

Remove saturated Kimwipe from Omnitray and incubate Omnitray with the device inside

at 37 °C, 5% CO2, for 12 hours.

b. Prior to cell loading, prepare two 5mm thick, 1 cm X 0.5 cm, sterile PDMS posts and

place in the Omnitray containing PMMA device approximately 70 cm apart. Dispense 50

μL suspended cell solutions into bottom compartment of the PMMA device through the

Appendix A 124

inlet ports. Dispense 80 µL of fresh supplemented media into the top compartment of the

device through the inlet ports. Remove saturated Kimwipe from Omnitray. Turn the

PMMA device over and place faced down on the two PDMS posts and incubate

Omnitray with the device inside at 37 °C, 5% CO2, for 12 hours.

9. After 12 hours, flush top and bottom compartments of devices with fresh media 2 times

to remove unattached cells.

10. Incubate at 37 °C, 5% CO2 with fresh media and keep devices in the up-right position for

remainder of culture period.

11. Replace media from both the top and bottom compartments every 24 h.


1. For LIVE/DEAD assay aspirate out old media from top and bottom compartments of

device and flush with serum-free media 3 times.

2. Prepare solution with 2μM Calcein AM, 4 μM ethidium homodimer -1 and Hoechst

33342 at (1:1000) dilution in serum-free media. Depending on the total volume a conical

tube or Eppendorf tube may be used to contain solution. Vortex solution for 10 to 15

sec using mini vortex mixer.

3. Dispense 80 µL and 50 µL of prepared staining solution into top and bottom

compartments, respectively, of the device.

4. Incubate at 37 °C, 5% CO2 for 30 min.

5. Flush top and bottom compartment with PBS 3 times. Begin here for actin and junction

protein staining.

6. After staining and flushing with PBS, fix cells with 4% PFA by loading solution through

the inlet ports of the top and bottom compartments and keep at room temperature for 20

min.

7. Flush the device with PBS 3 times to remove all PFA from the device. Gels may be

extracted from the device and imaged for LIVE/DEAD assay (See below for Imaging

protocol).

8. Permeabilize cells by flushing top and bottom compartments with 0.1% Triton X-100 and

keeping at room temperature for 5 min, this step not necessary for epithelial barrier

function staining. Flush compartments with PBS 3 times immediately after 5 min

permeabilization period.

9. Block sample with blocking buffer (3% bovine serum albumin (BSA) with 0.1% Tween 20

in PBS) by dispensing solutions into top and bottom compartment of the device and

incubating for 3 h at 37 °C.

10. Prepare staining solution:

a. For ZO-1 labelling prepare rabbit anti-ZO-1 pAb (1:200 dilution in blocking buffer).

b. For α-SMA labelling prepare with mouse anti-α- SMA mAb (1:200 dilution in blocking

buffer).

c. For co-culture staining, anti-bodies may be mixed in their respective dilutions in the

same blocking buffer solution.

11. Dispense staining solution into top and bottom and leave device in a Omnitray in the

fridge for 48 h at 4oC.

12. After labelling, flush compartments with PBS 3 times.

Appendix A 125

13. Prepare solution with:

a. For Calu-3 monoculture samples labelled with Rabbit anti-ZO-1 pAb only, prepare

AlexaFluor 568 goat anti-rabbit IgG (1:200 dilution in blocking buffer) for conjugation

with ZO-1 and Hoechst 33342 (1:1000 dilution) to stain the nuclei. To co-stain for

F-actin, add ActinGreen 488 (2 drops per ml of blocking buffer) into solution.

b. For Calu-3 monoculture samples labelled with mouse anti-α- SMA mAb only, prepare

AlexaFluor 568 goat anti-mouse IgG (1:200 dilution in blocking buffer) for conjugation

with α-SMA and Hoechst 33342 (1:1000 dilution) to stain the nuclei. To co-stain for

F-actin, add Alexa Fluor 488 phalloidin (2.5% (v/v) in the solution.

c. For co-culture staining, fluorescent stains may be mixed in their respective dilutions in

the same blocking buffer solution

14. Incubate sample at 37 °C, 5% CO2 for 2 hours. Flush compartments with PBS 3 times to

remove excess fluorescent labelling solution.

Sample preparation for Imaging

1. To extract samples from the device. Detach top layer from the bottom two layers of

PMMA using a single edge steel blade.

2. Lodge blade between the top and the middle layer at one corner of the device. While the

blade is lodged in, move along the four edges of the device to slowly open the

PMMA-PMMA bonds between the top and middle layer.

3. Move the blade across the device while still lodged between the PMMA layers to detach

the top layer.

4. Once the top layer is removed, detach the edges of the hydrogel using the tip of the

sharp scalpel.

5. Gently rinse the hydrogels using a pipette with PBS until the hydrogels completely

detach from the device

6. Place hydrogel on a 1.2 mm thick 75 X 25 mm glass microscope slide for imaging.

Appendix A 126

A6.0 Cell Adhesion Data

Table A6.1 Area covered by Calu-3 cell culture on Day 2

System # for each

case

Col-I (10.31µg/µL)

Col-I (6mg/L) + Matrigel (3mg/L)

Col-I (3µg/µL) + Matrigel (6µg/µL)

Matrigel (9µg/µL)

Area covered by Calu-3 cell culture

on Day 2 (%)

1 45 83 92 40

2 65 86 96 36

3 72 76 89 52

4 76 79 87 31

Table A6.2 Area covered by Calu-3 cell culture on Day 7

System # for each

case




Matrigel (9µg/µL)

Area covered by Calu-3 cell culture

on Day 7 (%)

1 13 16 92 34

2 18 18 85 46

3 9 20 83 62

4 15 12 87 39

Table A6.3 HBSMCs adhered per mm²of hydrogel surface on Day 2

System # for each

case




HBSMCs adhered per mm²of hydrogel

surface on Day 2

1 1047 957 1304

2 1743 1583 2045

3 1233 1628 1456

4 1341 1681 1903

Appendix A 127

Table A6.4 HBSMCs adhered per mm²of hydrogel surface on Day 7

System # for each

case




HBSMCs adhered per mm²of hydrogel

surface on Day 7

1 5123 4282 4978

2 4696 4978 4346

3 3467 4923 4346

4 4967 4178 4123

Table A6.4 Area covered by Calu-3 cell culture seeded at various cell seeding densities.

1000 cells/mL 1500 cells/mL 3000 cells/mL

Area covered by Calu-3 cell

culture seeded at

various cell seeding

densities (%)

Day 3 22 16 18 25 35 40 32 26 67 55 68 52

Day 5 24 27 30 31 47 36 52 55 57 82 63 75

Day 8 37 35 34 29 65 79 73 58 69 81 55 78

Day 14 41 46 43 36 73 64 85 87 97 89 79 92

Day 21 62 73 67 62 94 91 99 96 97 99 95 93

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