combining fabrication and surface ......combining fabrication and surface modification techniques to...

COMBINING FABRICATION AND SURFACE MODIFICATION TECHNIQUES

TO DEVELOP CELL-LADEN MICROFLUIDIC DEVICES

A Thesis

Submitted to the Faculty

of

Drexel University

by

Qudus Hamid

in partial fulfillment of the

requirements for the degree

of

Doctor of Philosophy

in

Mechanical Engineering and Mechanics

June 2014

iii

DEDICATED TO

To my superb family:

My wife, Sreylark Som

My son, Dylan Sot Hamid

My Parents, Keith Ivelaw Hamid and Bissoondai Hamid

My Brothers, Emron Hamid, Aneil Hamid, Murvin Hamid, Babak Hamid, and Abbas Hamid

My Sister, Amanda Frenceska Hamid

My Nephew, Nicholas Ivelaw Hamid

My Nieces, Carrey Hamid, Priya Hamid, Cassie Hamid, and Kelly Hamid

…..with love and admiration

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“Two little mice fell in a bucket of cream. The first mouse quickly gave up and drowned. The second mouse, wouldn't quit. He struggled so hard that eventually he churned that cream into

butter and crawled out. Gentlemen, as of this moment, I am that second mouse.”

-Frank Abagnale Sr.

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ACKNOWLEDGEMENTS

My sincerely thanks and appreciation goes to my advisor and mentor, Dr. Wei Sun, for his

unconditional encouragement, supervision, and leadership throughout my academic career at

Drexel University. His support and guidance has matured me into the researcher that I am today.

I would thank the members of my committee, Dr. MinJun Kim, Dr. Leslie Lamberson, Dr.

Jack Zhou, Dr. Binil Starly, and Dr. Yinghui Zhong for their valuable comments and time spent on

improve the contents of this thesis.

For supporting my research activities during my doctoral studies, I would like to

acknowledge; the National Science Foundation for the East Asian and Pacific Summer Institute

Fellowship to the People’s Republic China: Grant No. 1209517, two Summer Institute Short

Courses Fellowship, and research grant NSF-CMMI-1030520, Johnson & Johnson – Advanced

Technologies & Regenerative Medicine, LLC (ATRM), and Drexel University’s Office of

Graduate Studies (OGS) Parental Accommodation Fellowship.

Most importantly, for the late-nights, debates, the creative questions, and laughs, I would

like to knowledge my colleagues, friends, and past and present Computer-aided Tissue Engineering

Laboratory (CATEL) and Biofabrication Laboratory (BFLab) members. Specifically, Steven K.

Leist, Paul S. Kim, U Kei Cheang, Chengyang Wang, Jessica Snyder, Yigong Liu, Shannon

Williams, Eric Tran, Stephan Tran, George Yan, Hoyeon Kim, Mishah Salman, DalHyung Kim,

Tom Meleey, Ryan Robinson, Kathleen Donahue, Lauren Shor, Kalyani Nair, Eda Yildirim,

Bobby Chang, Teck-Kah Lim, Mark Timmer, and Taz Kwok.

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TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................................ X

LIST OF FIGURES ..................................................................................................................... XI

ABSTRACT ............................................................................................................................... XVI

CHAPTER 1: INTRODUCTION .............................................................................................. 1

1.1 Need for Cell-laden Microfluidics ............................................................................................. 1

1.2 Challenges and Current Fabrication Approaches ..................................................................... 7

1.3 Advantages of Cell-laden Microfluidic Chips and Maskless Fabrication ............................... 18

1.4 Research Objectives and Approach ......................................................................................... 21

1.5 Thesis Outline .......................................................................................................................... 22

CHAPTER 2: SURFACE MODIFICATION OF SU-8 FOR ENHANCED CELL

ATTACHMENT AND PROLIFERATION .............................................................................. 25

2.1. An Inspection of Surface Modification .................................................................................... 25

2.2. The Development of a Bare SU-8 Substrate ............................................................................ 29

2.3. Water Contact Angle Investigations ........................................................................................ 34

2.4. Topological Analysis ............................................................................................................... 36

2.5. X-Ray Photoelectron Spectroscopy (XPS) Analysis ................................................................ 39

2.6. Biological Investigations ......................................................................................................... 45

2.7. Interpretations ......................................................................................................................... 48

CHAPTER 3: UTILIZATION OF A DYNAMIC DIGITAL MICRO-MIRRORING

SYSTEM WITH A MULTI-NOZZLE BIOLOGICS DEPOSITION SYSTEM TO

FABRICATE CELL-LADEN MICROFLUIDICS ................................................................... 50

3.1. Applications of a Digital Micro-mirroring System .................................................................. 50

3.2. Digital Micro-mirroring System .............................................................................................. 53

3.3. Multi-nozzle Biologics Deposition System ............................................................................... 57

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3.4. Microfluidic Chip Fabrication and Characterization Protocols ............................................. 59

3.5. Cell Proliferation, Cytotoxicity Analysis, and Cell Morphology ............................................. 63

3.6. Cell Printing and Structural Integrity ..................................................................................... 67

3.7. Conclusions ............................................................................................................................. 70

CHAPTER 4: INTRODUCTION OF A FREEFORM MICRO-PLASMA SYSTEM FOR

THE DEVELOPMENT OF A THREE-DIMENSIONAL CELL-LADEN MICROFLUIDIC

CHIP OF IN VITRO DRUG METABOLISM DETECTION .................................................. 71

4.1 A Synopsis of Cell-laden Microfluidic Chips ........................................................................... 71

4.2 System Overview ...................................................................................................................... 73

4.3 Development of Three-dimensional Interconnected Microfluidic Chips ................................. 76

4.4 Sterilization, Plasma Treatment, and Cell Printing ................................................................ 81

4.5 Cytotoxicity Analysis and Cell Interactions............................................................................. 83

4.6 Drug Metabolism, Cell Morphology and Structural Integrity ................................................. 88

4.7 Fluid Dynamics Computational Analysis ................................................................................ 91

4.8 Limitations and Challenges ..................................................................................................... 94

CHAPTER 5: INTEGRATING THE MULTI-NOZZLE BIOLOGICS DEPOSITION

AND MICRO-PLASMA SYSTEMS WITH A FREEFORM ULTRA-VIOLET HEAD AND

A PHOTO-POLYMER MATERIAL DELIVERY SYSTEM TO INVESTIGATE CO-

CULTURE OF CANCER CELLS IN A MICROFLUIDIC ENVIRONMENT .................... 97

5.1 Feasibility of Testing Protocols, Availability, and Ethical Concerns ...................................... 97

5.2 System Integration Analysis ................................................................................................... 101

5.3 Manufacturing Methods ........................................................................................................ 106

5.4 Biological Characterizations ................................................................................................. 109

5.5 System Characterization ........................................................................................................ 110

5.6 Cell integration, Proliferation, and Morphological Investigations ....................................... 118

5.7 Limitations and Challenges ................................................................................................... 123

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CHAPTER 6: CONCLUSIONS AND RECOMMENDATIONS ....................................... 124

6.1 Summary of the Research ...................................................................................................... 124

6.2 Research Contributions ......................................................................................................... 127

6.3 Future Research Recommendations ...................................................................................... 129

LIST OF REFERENCES .......................................................................................................... 132

VITA ........................................................................................................................................... 160

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LIST OF TABLES

Table 1-1. Bio-modeling design requirements and possible solutions(B. Starly, 2006). ............... 11

Table 2-1. Water Contact Angle Measurement. ............................................................................ 36

Table 2-2. Quantitative Analysis of Each Surface Treatment........................................................ 37

Table 2-3. Atomic Elemental Composition of Each Treated Surface ............................................ 44

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LIST OF FIGURES

Figure 1-1. Adopted from Esch et al, to illustrate a sample of a body-on-a-chip device. This figure presents a schematic of how microfluidic cell culture systems can be used in conjunction with other in vitro cell-based assays, mathematical models, and in vivo experiments to enhance the drug development process and improve toxicity estimations for environmental contaminants (Esch et al., 2011). ............................................................................................................................................... 6

Figure 1-2. Three-dimensional reconstruction roadmap (W. Sun & Lal, 2002b). ......................... 14

Figure 1-3. Screenshot of a 3D-R process on Materialise Mimics. ............................................... 15

Figure 1-4. Adopted from Nguyen et al, this schematic illustrates a drug delivery microfluidic device implanted onto an eye (N.-T. Nguyen et al., 2013). ........................................................... 18

Figure 2-1. A schematic of the digital micro-mirroring microfabrication system. ........................ 30

Figure 2-2. Schematic of the multi-nozzle biologics deposition system (R. Chang, Sun, W.,, 2009). ....................................................................................................................................................... 31

Figure 2-3. (A) Model of the PDMS enclosure, (B) micro-channel fabricated within the bottom enclosure of the chip. ..................................................................................................................... 32

Figure 2-4. The multi-nozzle biologics deposition system printing cells within the channels of the chip................................................................................................................................................. 33

Figure 2-5. Side-view images showing water droplet illustrating the water contact angle on: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces. ............ 35

Figure 2-6. Three-dimensional profile of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces. .................................................................................................. 38

Figure 2-7. Line profile of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces. ...................................................................................................................... 39

Figure 2-8. XPS survey spectra of untreated surfaces. .................................................................. 40

Figure 2-9. XPS survey spectra of the gelatin treated surface. ...................................................... 41

Figure 2-10. XPS survey spectra of the plasma treated surface. .................................................... 41

Figure 2-11. XPS survey spectra of the sulfuric acid treated surface. ........................................... 42

Figure 2-12. Detailed XPS spectra of carbon 1s for the untreated surface. ................................... 42

Figure 2-13. Detailed XPS spectra of carbon 1s for the gelatin treated surface. ........................... 43

Figure 2-14. Detailed XPS spectra of carbon 1s for the plasma treated surface. ........................... 43

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Figure 2-15. Detailed XPS spectra of carbon 1s for the sulfuric acid treated surface. .................. 44

Figure 2-16. Cell proliferation study of untreated, gelatin treated, plasma treated, and sulfuric acid treated surfaces. ............................................................................................................................. 46

Figure 2-17. Cell morphology of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces. ......................................................................................................... 48

Figure 3-1. Applications of the dynamic digital micro-mirroring microfabrication system .......... 52

Figure 3-2. Structure of the digital micro-mirroring microfabrication system. ............................. 53

Figure 3-3. Digital micro-mirroring microfabrication system. ...................................................... 54

Figure 3-4. Illustration of light reflection on the digital mirrors.................................................... 55

Figure 3-5. (A) The digital micro-mirroring transmission spectrum, (B) The Ultraviolet source relative intensity range. .................................................................................................................. 56

Figure 3-6. Structure of the photolithographic substrate alignment system. ................................. 57

Figure 3-7. An image of the major components of the Multi-nozzle Biologics Deposition System. ....................................................................................................................................................... 58

Figure 3-8. Pneumatic micro-valve nozzle for the multi-nozzle biologics deposition system. ..... 59

Figure 3-9. 14 day cell proliferation study of treated and untreated open and closed microfluidic chips. .............................................................................................................................................. 65

Figure 3-10. (A) A fluorescence image, taken at 14 days after cells were seeded into the microfluidic chip showing live cell stained green and dead cells stained red. (B) A confocal image, taken 24 hours after cells were seeded into the microfluidic chips showing the nuclei (stain bright green) and the cytoplasm (stain green) of the cells in the channel. (C) An SEM image, showing an in-depth view of the cell morphology within the channels. ........................................................... 67

Figure 3-11. The effects of conventional and cell printing seeding methods on cell proliferation within the microfluidic chips. ........................................................................................................ 68

Figure 3-12. (A) A schematic of the microchannels on the microfluidic chips. (B) An image of the left side of a microchannels on the microfluidic chip showing the cells (labeled with the arrows) within the channel and channel’s uniformity. (C) An image of the center of a microchannels on the microfluidic chip showing the cells (labeled with the arrows) within the channel and channel’s uniformity. (D) An image of the right side of a microchannels on the microfluidic chip showing the cells (labeled with the arrows) within the channel and channel’s uniformity. ............................... 69

Figure 4-1. A flow chart of the micro-plasma system. .................................................................. 74

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Figure 4-2. A schematic showing the cross-section of the micro-plasma nozzle treating the surface of a substrate. The major components of the nozzle are shown along with a photo of a treated sample is illustrated to demonstrate the effects of the micro-plasma nozzle. ................................ 75

Figure 4-3. A Model of the PDMS enclosure of the microfluidic chip. ........................................ 77

Figure 4-4. (A) A schematic of the fabrication and assembly of the cell-laden microfluidic chip. (B) A schematic illustrating the microchannel orientation of the first and second layers of the chip along with a schematic of the two layers overlapping each other for the 300 µm chip, the black bars represents the channel walls and the white bars are the channels. (C) A schematic illustrating the microchannel orientation of the first and second layers of the chip along with a schematic of the two layers overlapping each other for the 300 µm chip, the black bars represents the channel walls and the white bars are the channels. (D) A schematic illustrating the microchannel orientation of the first and second layers of the chip along with a schematic of the two layers overlapping each other for the 300 µm chip, the black bars represents the channel walls and the white bars are the channels. ........................................................................................................................................ 80

Figure 4-5. (A) A photo of the biologic deposition nozzle printing cells into the channels of the chip. (B) A photo of a fully fabricated chip, complete with enclosure, inlet and outlet ports (white), and internal features. (C) A photo of the incubation period of the chips where the syringe pump perfuse culture medium through the chips. .................................................................................... 82

Figure 4-6. (A1, B1, C1) Optical images of a pore of the interconnected chips. The dashed lines highlights the channel walls, the arrow points at cells within the channels. A1 is an optical image of a 300 µm pore chip, B1 is an optical image of a 500 µm pore chip, and C1 is an optical image of a 700 µm pore chip. (A2, B2, C2) are fluorescence images of the live cells stained green with the live dead assay. These images highlight the live cell’s orientation and uniformity within the channels of each chip. A1 is a fluorescence image of a channel in the 300 µm pore chip, B2 is a fluorescence image of a channel in the 500 µm pore chip, and C2 is a fluorescence image of a channel in the 700 µm pore chip. ................................................................................................... 85

Figure 4-7. Results of the 14 day proliferation investigation of the 300 µm, 500 µm, and 700 µm microfluidic chips. ......................................................................................................................... 87

Figure 4-8. Results of the EFC Drug concentration in the 300 µm, 500 µm, and 700 µm chips over a 12 hours period. ........................................................................................................................... 89

Figure 4-9. (A) A SEM images showing the cross-sectional of a microfluidic chip. This image illustrates the channel’s formation and structural integrity. Each chip showcases the same formation and structural integrity with their corresponding varying channel width. (B) A SEM image showing the morphology and attachment of the MDA-MB-231 cells within the microchannel of the chip. ..................................................................................................................................... 90

Figure 4-10. COMSOL Multiphysics simulations illustrating the fluid flow within the 300 μm, 500 μm, and 700 μm microfluidic chips. (A1) is a streamline simulation of the fluid flow within the 300 μm interconnected channels. (A2) is a velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout the 300 μm microfluidic chip. (A3) is a close-up of the velocity gradient at one of the interconnected pore within the 300 μm microfluidic chip. (B1) is a streamline simulation of the fluid flow within the 500 μm interconnected channels. (B2) is a velocity gradient

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showing the magnitude, direction, and fluid flow type that exist throughout the 500 μm microfluidic chip. (B3) is a close-up of the velocity gradient at one of the interconnected pore within the 500 µm microfluidic chip. (C1) is a streamline simulation of the fluid flow within the 700 μm interconnected channels. (C2) is a velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout the 700 μm microfluidic chip. (C3) is a close-up of the velocity gradient at one of the interconnected pore within the 700 µm microfluidic chip. Color scale bar unit: µm/s ....................................................................................................................................... 93

Figure 5-1. Adopted from Junttila et al, this schematic illustrates heterogeneity of a cancer model (Junttila & de Sauvage, 2013) ........................................................................................................ 99

Figure 5-2. (left) an image of the integrated fabrication system, (right) close-up of the four fabrication head respectively labeled. .......................................................................................... 102

Figure 5-3. (A) image of the three-dimensional spatial control system with its major components labeled, (B) an image of the photo-polymer head with its major components labeled, (C) a cross-sectional schematic of the localized micro-plasma head with its major components labeled, (D) a cross-sectional schematic of the biologics head showing its major components, (E) An image of the freeform ultra-violet micro-nozzle with its major components labeled. ................................ 105

Figure 5-4. Flow chart of the integrate system with each of its five major components outlined with color-coded dashed lines. ............................................................................................................. 106

Figure 5-5. (A) a schematic illustrating the fabrication steps of developing the cell-laden microfluidic chip, (B) a model of the PDMS enclosure, (C) an image of the fabricated microchannels within the slot of the PDMS enclosure, (D) an image of the completed cell-laden microfluidic chip with the lid and its inlet and outlet ports. ........................................................ 108

Figure 5-6. Percentage of live cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008). ................................................................................................... 113

Figure 5-7. Percentage of injured cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008). ................................................................................................... 114

Figure 5-8. Percentage of dead cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008). ................................................................................................... 114

Figure 5-9. Surface plot for the percentage of live cells as a function of process parameters (Kalyani Nair, 2008). .................................................................................................................................. 116

Figure 5-10. Surface plot for the percentage of dead cells as a function of process parameters (Kalyani Nair, 2008). ................................................................................................................... 116

Figure 5-11. Surface plot for the percentage of injured cells as a function of process parameters (Kalyani Nair, 2008). ................................................................................................................... 117

Figure 5-12. (A) SEM image showing the uniformity of the fabricated microchannels, (B) SEM image showing the end of the microchannel in which the direction changes from a horizontal channel to a vertical channel then back to a horizontal channel. ................................................. 118

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Figure 5-13. (A) fluorescence image showing cell distribution and integration of the MDA-MD-231 cells (red, Qtracker® 625) and the HepG2 cells (green, Qtracker® 525) within the microchannels, (B) a phase-contrast image of the cells in the microchannel, (C) quantitative results of the cell distribution of the MDA-MB-231 and HepG2 cell lines within the microfluidic chip. ..................................................................................................................................................... 119

Figure 5-14. Results of the 21 days cell proliferation study of the MDA-MB-231 cell-laden chip (control 1), HepG2 (control 2) cell-laden chip, and the co-culture (both MDA-MB-231 and HepG2 cell lines) cell-laden chip. ............................................................................................................ 120

Figure 5-15. (A) SEM image showing an overview of the cell distribution within the microchannel, (B) SEM image showing a close-up of the cells within microchannel, the MDA-MB-231 and HepG2 cells are labeled, (C) SEM image showing the morphology of a MDA-MB-231 cell, (D) SEM image showing the morphology of a HepG2 cell. .............................................................. 122

Figure 6-1. Flow chart illustrating the fabrication process of a cell-laden microfluidic chip using the integrative fabrication process. .............................................................................................. 126

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ABSTRACT

Combining Fabrication and Surface Modification Techniques to Develop Cell-laden

Microfluidic Devices

Qudus Hamid Wei Sun, Ph.D.

Micro-Electro-Mechanical Systems (MEMS) technologies illustrate the potential for many

applications in the field of tissue engineering, regenerative medicine, and life sciences. The

fabrication of tissue models integrates the multidisciplinary field of life sciences and engineering.

Presently, monolayer cell cultures are frequently used to investigate potential anti-cancer agents.

These monolayer cultures give limited feedback on the effects of the micro-environment. A micro-

environment, which mimics that of the target tissue, will eliminate the limitations of the traditional

mainstays of tissue research. The fabrication of such micro-environment requires a thorough

investigation of the actual target organ and/or tissue. Microfabrication techniques are utilized to

develop microfluidic channels for continuous nutrition supply to cells inside a micro-environment.

The ability of cells to build tissues and maintain tissue-specific functions depends on the interaction

between cells and the extracellular matrix (ECM). Three-dimensional tissue platforms are rapidly

becoming the method of choice for quantification of the heterogeneity of cell populations for many

diagnostic and drug therapy applications. Microfluidic sensors and the integration of sensors with

microfluidic systems are often described as miniature versions of their macro-scale counterparts.

This technology presents unique advantages for handling costly and difficult-to-obtain samples and

reagents as a typical system requires between 100 nL to 10µL of working fluid. The fabrication of

a fully functional cell-based biosensor utilizes both biological patterning and microfabrication

techniques. SU-8 is a popular photosensitive epoxy-based polymer in MEMS. The patterning of

bare SU-8 alone does not provide the appropriate ECM necessary to develop microsystems for

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biological applications. Manipulating the chemical composition of SU-8 will enhance the

biological compatibility, giving the fabricated constructs the appropriate ECM needed to promote

a functional tissue array. The objective of this research is to investigate the integration of maskless

fabrication, direct cell deposition, and surface modification techniques to engineer cell-laden

microfluidics. This thesis presents advances in additive manufacturing techniques, the utilization

of plasma chemistry to enhance surface functionalization, and manipulation of photo-

polymerization to investigate new approaches to assemble cell-laden microfluidics.

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CHAPTER 1: INTRODUCTION

1.1 Need for Cell-laden Microfluidics

Humans have occupied the earth for over one million years. The human body has evolved

throughout time to sustain life(Cavalli-Sforza, Piazza, Menozzi, & Mountain, 1988). Researchers

are constantly conducting analysis to understand human anatomy and physiological needs. A better

understanding of the human body, along with its tissues and organs, will enable scientists to develop

tissue substitutes to replace damaged and/or failed tissue organs(Zubal et al., 1994). The need for

replacement organs and tissue substitutes are on the rise (Ringeisen et al., 2013). Presently, there

are not sufficient amount of tissue replacements for failed or damaged organs, due to the lack of

donors. In the United States alone, there are over twenty million patients per year that suffer from

some form of tissue and/or organ related maladies, and are awaiting a replacement. The financial

cost of health care for these patients has been estimated to be over $400 billion annually (Klein et

al., 2010; Merion, 2010). Cell-laden microfluidic constructs are one of several promising

applications to address this issue. These constructs are fabricated from a variety of science and

engineering disciplines to create the optimum tissue replacement (in terms of the targeted

functionality). Additionally, these constructs play a vital role as pre-formed extracellular matrices

onto which cells can readily attach, rapidly multiply and form new tissue (Zein, Hutmacher, Tan,

& Teoh, 2002; Zeltinger, Sherwood, Graham, Mueller, & Griffith, 2001). Recently, the U.S.

government funded an excess of $24 million to study the feasibility and development of a functional

body-on-a-chip (Hughes, 2010).

Human physiological systems are modeled in complexity of scale (N.-T. Nguyen, Shaegh,

Kashaninejad, & Phan, 2013). It is agreed that the body comprises of tissues and organs that

function sequentially to sustain the viability and function of a person(Butcher, Berg, & Kunkel,

2004; Lambert, Gibson, & Noakes, 2005). Organs in the body utilize the nervous and endocrine

2

system to communicate. On the other hand, organs and tissues need a constant supply of essential

nutrients to operate on a daily basis. The circulatory system is the pipeline in which these nutrients

are delivered (Silverthorn, Ober, Garrison, Silverthorn, & Johnson, 2009). These

systems/functions coupled together are classed as a macro-scale system(s). At the micro-scale

level, each tissue and organ has its own unique architecture, function(s), different cell types, and

cell-cell interactions (P. X. Ma & Zhang, 2001). To understand the function of each organ and

tissue, scientists must examine structural integrity and functionality at the micro-scale level.

Information gathered at the micro-scale level can then be utilized to develop functional organs at

the macro-scale (Springer, 1990). Since scientists cannot build a fully functional tissue or organ at

a macro-scale (as yet); many researchers has decided that understanding an organ or tissue’s

function at the micro-scale level may lead to economical investigation, fewer errors in qualitative

and quantitative assessments, and most importantly; replicating the organ or tissue’s function

without fabricating the exact architecture of the actual tissue/organ (S. N. Bhatia & Chen, 1999;

Chung et al., 2009; Crevillén, Ávila, Pumera, González, & Escarpa, 2007). Once the tissue’s

function is fully analyzed and scientists has deemed it on par with that of the actual tissue, this

model will then serve as a building block which will be utilized for the development of a tissue or

organ on the macro-scale level.

It is essential that scientists replicate the exact function of the targeted tissue, organ or

disease in question. For the development of tissue and disease models; mechanical, chemical, and

biological cues of the target organ are carefully examined (B. M. Baker & Chen, 2012; S. N. Bhatia

& Chen, 1999; Causa, Netti, & Ambrosio, 2007) (Brandl, Sommer, & Goepferich, 2007; M. P.

Lutolf & J. A. Hubbell, 2005). Due to conventional manufacturing limitations, it is extremely

difficult to mimic the exact micro-architecture of the targeted tissue/organ (Leong, Cheah, & Chua,

2003). Tissue and disease models are developed by fabricating a platform that closely mimics the

3

functionality of the targeted organ. Quite often, the micro-architecture of these models are different

than that of target organ. Since one of the objectives is to develop a model that mimics the targeted

organ’s function; the design of the tissue model is developed to provide the appropriate mechanical,

chemical, and biological cues necessary to accomplish this objective (B. Starly, 2006). The

development of tissue and disease models enable researchers to; 1) understand and mimic tissue

function in vitro (Courtney, Sacks, Stankus, Guan, & Wagner, 2006; Linda G Griffith & Swartz,

2006; Tortelli & Cancedda, 2009), 2) investigate pharmaceutical products (Jasch et al., 2009; L.

Kang, Chung, Langer, & Khademhosseini, 2008), 3) develop building blocks for the assembly of

functional tissue organs (Dietmar W Hutmacher, Michael Sittinger, & Makarand V Risbud, 2004;

Jakab et al., 2010; Vladimir Mironov et al., 2009), 4) understand cell integration and migration

(Chung et al., 2009; H. Lu et al., 2004; Meyvantsson & Beebe, 2008), 5) develop biosensors for

counter-terrorism (D. Lu, Cagan, Munoz, Tangkuaram, & Wang, 2006; Nambayah & Quickenden,

2004; J. Wang, Thongngamdee, & Lu, 2006), and 6) develop sensors and alarms for domestic use

(Pumera, Merkoçi, & Alegret, 2006). Single-cell-based arrays do not capture multi-organ/multi-

cell interactions and limit the capabilities of the arrays (Esch, King, & Shuler, 2011). Tissue models

with a heterogeneous architecture are much more effective. Freeform fabrication techniques have

enabled researchers to develop tissue models with multiple cells, biologics, and biological

materials. The heterogeneity of these micro-systems have paved the way to develop complex tissue

constructs (S. Bhatia, Balis, Yarmush, & Toner, 1999; Khetani, Szulgit, Del Rio, Barlow, & Bhatia,

2004; Takayama, Taniguchi, & Okano, 2007).

When a drug is taken; it is absorbed, distributed, metabolized, and eventually, eliminated

(Esch et al., 2011; N.-T. Nguyen et al., 2013). Micro-scale cell-laden tissue arrays enable

researchers to develop economical testing platforms to investigate these processes of a targeted

drug. These micro-tissue arrays allow for minimal amount of testing material to produce

4

quantitative and qualitative results. In vitro cell-laden microfluidic systems can give the first

indication of the toxicity and efficacy of a compound (Bhushan et al., 2013; Esch et al., 2011). The

micro-systems are beneficial in terms that many investigations can be conducted simultaneously at

a fraction of the cost of clinical investigations. Apart from the cost-effectiveness of these micro-

arrays; drug investigations can be initiated faster since these platforms does not require government

approval. Since these testing platforms can potentially give the same results as those of animal

investigations; the use of micro-tissue arrays will significant reduce the need for animals for clinical

trials (Linda G Griffith & Naughton, 2002; Mancinelli, Cronin, & Sadée, 2000). For micro-

systems to be successful, they must mimic the function(s) of that being targeted by the

pharmaceutical product. Many pharmaceutical companies are developing micro-systems that

mimic the functionality of the liver and target organs to allow for faster and cheaper investigation

of their products (L. Kang et al., 2008). These biological micro-constructs will lead to the

development of better pharmaceutical products which will aid in prolonging human life.

In addition to pharmaceutical platforms, there is a significant need for biological sensors

for both counter-terrorism (D. Lu et al., 2006; Nambayah & Quickenden, 2004; J. Wang et al.,

2006) and domestic uses (Pumera et al., 2006). Over the last decade, there has only been 6 approved

biologic license applications in a field where it is estimated that about 1 of 10,000 compounds has

been successfully tested (Hughes, 2010). Consumer products such as the glucose monitoring

system (Lebel et al., 1996) and pregnancy tests (Chard, 1992) are widely available and used

throughout the world. There is a huge market for consumer products that allows consumers to

conduct private investigations in the privacy of their homes (Bogue, 2007; Eloy & FEatuREs,

2010). Additionally, government agencies are working effort-lessly to develop new sensor

products to protect their respective borders and citizens from terror. Biological sensors are small

and have the potential to detect the smallest trace of toxic chemicals. The development of these

5

sensors will enable armed forces to prevent chemical and biological terroristic activities (D. Lu et

al., 2006; Nambayah & Quickenden, 2004; J. Wang et al., 2006).

The integration and interconnectivity of micro-systems can lead to the development of a

‘body-on-a-chip’ platform (Perozziello, Bundgaard, & Geschke, 2008). Body-on-a-chip platforms

are where unique micro-systems are ‘wired’ together to replicate the function(s) of the human body.

Presently, pharmaceutical investigations on micro-tissue arrays are very limited. With the

development of a body-on-a-chip platform, more extensive investigations are plausible. Body-on-

a-chip devices are utilized to improve the predictive power of in vitro screening tools (Esch et al.,

2011; Sung, Kam, & Shuler, 2010). Body-on-a-chip devices are not developed to make

replacement organs, instead, they are developed to replicate the targeted organ’s function to allow

for investigations of therapeutic and toxic effects (M. Baker, 2011). Body-on-a-chip devices are a

fairly new set of micro-systems and the potential of developing platforms other than for

pharmaceutical investigations are feasible. Figure 1-1 presents a schematic of a sample of a body-

on-a-chip device (Esch et al., 2011).

6

Figure 1-1. Adopted from Esch et al, to illustrate a sample of a body-on-a-chip device. This figure presents a schematic of how microfluidic cell culture systems can be used in conjunction with other in vitro cell-based assays, mathematical models, and in vivo experiments to enhance the drug development process and improve toxicity estimations for environmental contaminants (Esch et al., 2011).

7

1.2 Challenges and Current Fabrication Approaches

One of the challenges in tissue engineering is the inability to replicate the cellular and

system complexities of natural tissues and organs (Linda G Griffith & Naughton, 2002; Ikada,

2006). The field of biomaterials continues to yield a wealth of tissue and cell specific materials that

reproduce native cues and support attachment, proliferation, and migration of tissue specific cell

types. Bio-chemical co-factors are then added to stimulate differentiation and three dimensional

tissue developments (M. Lutolf & J. Hubbell, 2005). However, at the micro-scale level; cells and

co-factors are almost always randomly seeded, and then randomly attach to the micro-architectures.

This random attachment results in poor replication of the anisotropic cellular microniche that is

integral to defining the tissue phenotype and cellular response to stimuli (Ziółkowska, Chudy,

Dybko, & Brzózka, 2011). At the meso-scale, researchers struggle to create sustainable vasculature

throughout in vitro tissues, and engineering the complex architecture and function of vessels and

micro-capillaries (Linda G Griffith & Naughton, 2002; Nerem & Seliktar, 2001). Finally, at the

macro-scale, the concept of engineering multiple tissues that interact through an endothelialized

circulatory system and respond in a unified manner has not been realized. Additionally, the

appropriate cues (chemical, mechanical, and biological) must be captured and reproduced within

the tissue construct (B. M. Baker & Chen, 2012; S. N. Bhatia & Chen, 1999; Brandl et al., 2007;

Causa et al., 2007; M. P. Lutolf & J. A. Hubbell, 2005).

Scientists are good at fabricating tissue constructs that geometrically mimic (mechanical

cues) the target tissue. However, the constructs often lack functional groups (chemical and

biological cues) needed to sustain adequate cell attachment, proliferation, and differentiation.

Presently, a global treatment surface modification is added after the manufacturing process. The

issue with global treatments is that it is limited to one treatment per construct. A localized treatment

will provide the potential for various treatments within a single construct (Eda D Yildirim,

8

Besunder, Guceri, Allen, & Sun, 2008). Apart from surface modification, spatial control of cells

is essential for the development of a fully functional tissue construct. Conventional cell seeding is

performed by using a pippettor and dispensing cells (in culture medium) onto the tissue construct

(Lauren Shor, 2009). This conventional process does not allow for uniform distribution of cells

and is limited to one cell type per sample. The integration of a three-dimensional cell printer with

the localized surface treatment will enable scientists to have spatial control and print cells

immediately after surface modification.

Bio-Manufacturing Techniques. Bio-manufacturing is completely different compared to

conventional manufacturing techniques. Conventional manufacturing techniques would utilize a

raw stock material of which mills, drills, and cutting tools would remove materials from the raw

stock material to obtain the desired model. Bio-manufacturing does not build tissue constructs with

the use of mills, drills, and cutters (W. Sun, Yan, Lin, & Spector, 2006; Weigel, Schinkel, &

Lendlein, 2006). Instead, tissue constructs are bio-manufactured either by; 1) layer-by-layer

fabrication (Tang, Wang, Podsiadlo, & Kotov, 2007; Y. N. Yan et al., 2003; Zein et al., 2002), 2)

solid freeform fabrication (Dietmar W Hutmacher et al., 2004; Yarlagadda, Chandrasekharan, &

Shyan, 2005), or 3) photolithography (Bryant, Cuy, Hauch, & Ratner, 2007; Dong, Yong, Liao,

Chan, & Ramakrishna, 2008; Zhang, Hutmacher, Chollet, Poo, & Burdet, 2005). Depending on the

complexity and desired material, the appropriate fabrication technique is chosen. Other important

factors that determine the appropriate manufacturing techniques are the porosity, interconnectivity,

and transport property for nutrients that would enable the ingrowth of new cells and cell-tissue

formation. Successful tissue arrays have incorporated extracellular matrices (ECM) within their

respective tissue constructs. ECMs provide instructions that direct cell attachment, proliferation,

differentiation, and the growth of new tissue. Tissue constructs must also have incorporated within

it; heterogeneous characteristics in the form of scaffold materials, a controlled spatial distribution

9

of growth factors, and an embedded micro-architectural vascularization for cellular nutrition,

movement, and chemo-taxis. Consideration of these multiple biological, biomechanical and

biochemical issues can be represented by a comprehensive ‘tissue informatics’ model (Wei Sun,

Starly, Nam, & Darling, 2005).

Layer-by-layer manufacturing utilizes CAD/CAM technologies to fabricate tissue

constructs. CAD software would develop the three-dimensional model of the targeted organ. The

three-dimensional model would then be converted to the appropriate file type and uploaded to the

bio-manufacturing device. Layer-by-layer manufacturing devices use ‘.stl’ files to fabricate the

tissue construct. The ‘.stl’ file divides the three-dimensional model into layers. The bio-

manufacturing device would fabricate each layer, one after another. Layers are usually fabricated

directly above the previous layer and at the end of the fabrication process; a three-dimensional

tissue construct is produced (B. Starly, Lau, Sun, Lau, & Bradbury, 2005).

Solid freeform (SFF) bio-manufacturing devices have the ability to move in three-

dimensional space. These devices usually consist of three or more motion arms that enable the

material delivery system to fabricate the desired tissue construct. Most of the tissue constructs

fabricated from SFF technologies are porous features mainly because the material delivery

component of these devices produces a filament or droplet feature from its nozzle or printing head.

CAD and CAM technologies are used with SFF devices. CAD is utilized in the development of

the tissue model, while CAM technologies are used to manufacture the tissue construct. Unlike

layer-by-layer manufacturing techniques, SFF does not require its CAM files to be in the ‘.stl’

format. All SFF bio-manufacturing devices use various CAM files. Data that is taken from the

CAM files are coordinates, mass, volume, and toolpath of the fabrication head.

Photolithographic bio-manufacturing uses the layer-by-layer manufacturing approach to

fabricate its tissue constructs. Materials used with this manufacturing technique are sensitive to

10

light: once the material is exposed to light, it would change from liquid to a solid. One approach

of fabricating tissue constructs with photosensitive material is to add a small volume of the

photosensitive material within ‘petri dish’, then light (type of light is determined by the material’s

sensitivity) is used to produce the desired layer. More material is added and the process is repeated

until the modeled tissue construct is fabricated. The light in this manufacturing process can be

stationary and the ‘mask’ with the desired pattern is placed between the light and the substrate. In

addition, the light can be placed on one or more motion arms where the motion arm(s) would move

the light in a defined toolpath that would allow for the fabrication of the desired layer. The second

approach is to create a mold with the photosensitive material using the layer-by-layer approach.

The mold will be utilized to produce the desire tissue construct. This method is used when the

desired tissue construct has to be fabricated with a specific material that is not photosensitive.

Bio-modeling and biomimetic design. Conventional three-dimensional tissue constructs

are designed with a preferred internal architecture, wherein porosity and material connectivity

provide the required structural integrity, mass transport, and comprehensive micro-environment for

cell and tissue growth. Literature surveys has shown that cell survival and proliferation within the

tissue constructs are dependent on oxygen, vital molecules, and the micro-architecture of the

scaffolds (Maquet, 2007; Wake, 1994). The complexity of tissue scaffolds requires novel

approaches and computational algorithms to match the desired criteria for internal architecture,

permeability, pore size, and connectivity. The dynamics of a tissue construct are governed by

structural and topological configuration defined by porosity, pore interconnectivity, tortuosity,

material permeability and diffusivity (Mikos et al., 1994; Rajeev A, 2000; Ratner, 1996). The

tortuosity characterizes the diffusion path length of fluid molecules through the scaffold, which

shapes the internal architecture of the construct and plays a major role in tissue growth and

proliferation (E. A. Botchwey, M. A. Dupree, S. R. Pollack, E. M. Levine, & C. T. Laurencin,

11

2003; Hrabe, Hrabetova, & Segeth, 2004; Shuler & Kargi, 2002; Turing, 1990; Zalc, Reyes, &

Iglesia, 2004). Many cells respond more favorably to a three-dimensional micro-environment

(compared to a two-dimensional micro-environment) with intricate intracellular architectures

where the cell’s morphological shape, behavior, and gene expression are richer, more robust, and

closer to in vivo responses (Abbott, 2003; Albrecht, Tsang, Sah, & Bhatia, 2005; Benya & Shaffer,

1982). Tissue substitutes are designed to be replicas of actual tissue organs. Researchers gather

information about the tissue organ by conducting a biological investigation of the targeted organ

and developing a bio-model. The bio-model presents biological, chemical, mechanical, physical,

and structural information of the tissue organ. Each bio-model can be developed independently

and later be combined. Design requirements and solutions that are investigated during the

development of a tissue construct are listed in Table 1-1(B. Starly, 2006).

Table 1-1. Bio-modeling design requirements and possible solutions(B. Starly, 2006).

Properties Design Requirements Possible Design Solutions Mechanical Construct structural integrity

Internal architectural stability Construct strength and stiffness

Biomaterial selection Internal architecture Porosity and pore distribution Fabrication method

Geometrical Anatomical fitting Construct external geometry

Manufacturing Process ability Process effect

Advanced manufacturing using SFF based techniques Process controlled algorithms using appropriate process planning instructions

Biological Cell loading, distribution, and nutrition Cell attachment and in growth Cell-tissue aggregation and formation

Biomaterial selection Preferred internal architecture and layout Pore size and interconnectivity Vasculature

12

Tissue reconstruction. Images captured with the CT or MRI devices are two-dimensional

images. Without the third dimension it becomes very difficult to examine and analyze the tissue(s).

In order to properly visualize the tissue as a whole, multiple views and slices are displayed at the

same time. This method is very difficult for researchers, doctors, and surgeons to employ (Chow

& Sommer, 2001; Marko, Leith, & Parsons, 1988; Vannier, Marsh, & Warren, 1984; Weibel &

Elias, 1967). With the aided help of CAD software, it is possible to reconstruct the slices of

information into a three-dimensional model. Three-dimensional reconstruction (3D-R) will enable

researchers to directly study the tissue by displaying the three-dimensional anatomical images or

models. This newly constructed feature can be manipulated in terms of orientation, wireframe

models, hidden wireframe models, surface/shaded models, and solid models with or without the

following: reflectivity, variable lighting, and transparency. With a 3D-R model, volumes and

surface conditions may be determined.

The process of constructing a three-dimensional model from a set of CT scans usually

begins with the isolation of an area of interest. To convert a set of two-dimensional scans to a three-

dimensional model, all surfaces must be bounded to make a closed structure and all edges within

the area of interest must be defined. The roadmap which defines the process of 3D-R is illustrated

in Figure 1-2. In addition to the roadmap, listed below are the following benefits and issues with

3D-R models (W. Sun & Lal, 2002b):

i. Visualization and understanding of the anatomical boundaries of structures in three

dimensions, particularly those that are hidden. For example, an individual nerve fiber from

within a buddle.

ii. Intensity measurement: Three-dimensional models are considerably better than two-

dimensional models because there is no section thickness artifact. Although the use of

13

three-dimensional models improves the quantification of intensity, it also introduces a new

set of problems.

iii. The localization of entities. For example, dopamine-containing neurons in the human mid-

brain (Woodward, 1983). Co-localization studies of antigens demonstrated by

immunocytochemical techniques also fall into this category.

iv. The analysis of the distribution of the components: this can be at a tissue level. For

example, lymphoma deposits in bone marrow trephines (J. R. Salisbury, Deverell, M.H.,

1994; J. R. Salisbury & Whimster, 1994) at a cellular level.

v. Spatial quantitation, such as object counting; tissue density histograms. For example, the

three-dimensional distribution of tissue/cell numbers in different regions of the brain

(Woodward, 1983); and determination of volumes occupied by reconstructed structures.

vi. The examination of the relationships between components, such as whether they are

connected or not. For example, neurons in the cortex (Rydmark, Jansson, Berthold, &

Gustavsson, 1992) or notochordal tissue in embryos (J.R. Salisbury, 1992; J. R. Salisbury,

Deverell, Cookson, & Whimster, 1993).

The reconstructed three-dimensional models yield novel views of patient anatomy while

retaining the image voxel intensities that can be used for volume rendering, volumetric

representation and three-dimensional image representation. These three-dimensional images lead

to the generation of anatomic models which are used for contour based generation and three-

dimensional shaded surface representation of CAD based medical models. The generation of the

three-dimensional models is not a simple task; several visualization or computation errors may arise

during the generation process. Whenever an issue arises during the reconstruction process, a

prototype model is developed with additive/constructive processes, as opposed to subtractive

14

processes. Many applications have developed in recent decades to utilize 3D-R models to aid in

medical diagnosis (Jan et al., 2006), surgical planning (Piatt, Starly, Sun, & Faerber, 2006),

biomedical implants (P. Evans, Starly, B., Sun, W.,, 2006), and tissue engineering construct

development (Hollister, Maddox, & Taboas, 2002; D. W. Hutmacher, M. Sittinger, & M. V.

Risbud, 2004). Figure 1-3 shows a screenshot of the reconstruction process in Materialise Mimics

software.

Figure 1-2. Three-dimensional reconstruction roadmap (W. Sun & Lal, 2002b).

15

Figure 1-3. Screenshot of a 3D-R process on Materialise Mimics.

Micro-organs, Tissue-on-a-chip, and Lab-on-a-chip. Testing of pharmaceuticals and

biological compounds in humans or animals is not always possible, at least not in the early stage.

Moreover, while in vivo animal studies can provide data more relevant to human responses, animal

tests are expensive, labor-intensive, and time consuming (R. Chang, Sun, W.,, 2009; Gonda, 2008).

Accordingly, sometimes decisions need to be made based on in vitro data. However, extrapolating

in vitro data (for example, cell culture data) to the in vivo relevant conditions is often difficult.

Although pharmacokinetic principles can be used to derive some conclusions, this approach has

limitations. For example, cells cultures under traditional assay conditions may not function in the

same ways as cells would in natural settings because the communication and interactions between

16

different tissues and organs are absent. In culture, cells are typically grown at the bottom of

chambers or wells. These systems may have unrealistically high liquid-to-cell ratios. Even if the

cells are grown on the micro-carrier beads, which more closely resemble physiological conditions,

they still may not mimic physiological conditions accurately enough to provide reliable data (R.

Chang, Sun, W.,, 2009). Recent advances of micro- and nano-technologies along with the

integration of various components into a single micro-device have led to the development of lab-

on-a-chip devices. Lab-on-a-chip drug delivery devices are designed and developed to investigate

conventional delivery methods in which a drug administered through the mouth, the skin,

transmucosal areas, inhalation or injection. The Lab-on-a-chip devices investigate various

processes of release, absorption, distribution and elimination of drugs (Benet, Kroetz, Sheiner,

Hardman, & Limbird, 1996; N.-T. Nguyen et al., 2013). Since many of these lab-on-a-chip devices

lack the heterogeneity to that of the target organ, quite often, these results gathered from these

device will have to be confirmed with clinical trials. An advanced heterogeneous lab-on-a-chip

device will reduce the utilization of animal studies.

Micro-robots have been utilized with lab-on-a-chip and tissue-on-a-chip platforms to

isolate cells and deliver a specific dose to targeted area (Gao et al., 2012; D. H. Kim, Kim, Julius,

& Kim, 2012). Micro-robots vary in design, some are developed with the use of micro-organism

and/or their motor skills while others are purely abiotic. The objective of these micro-robots are to

one day be utilized to deliver a specific dose to a targeted area(s) in the body. Tissue-on-a-chip

platforms are devices currently used to characterize the capabilities of these micro-robots. As

previously mentioned, tissue-on-a-chip platforms offer the potential of modeling the critical tissues

and organ’s function of the body onto a microscopic platform. Although these platforms are very

advantageous, most tissue-on-a-chip devices are single cell platform which does not accurately

model the entire functionality of the target tissue/organ. Single cell tissue-on-a-chip platforms have

17

been developed to mimic a single target function of the tissue under investigation. Coupling and

integration tissue-on-chip platforms have led to the development of body-on-a-chip platform where

multiple functionalities of various organs and tissues can be investigated.

The lack of heterogeneous mimicry reduces the investigative potential of these microfluidic

chips. Many researchers have made the advances in developing heterogeneous microfluidic chips.

Three-dimensional tissue cultures in microfluidic chips have paved the way for the development of

heterogeneous microfluidic chips. To fully understand how tissues form and function, as well as

their pathophysiology, it is crucial to study how cells and tissues behave as parts of whole living

organs that are composed of multiple, tightly opposed tissue types that are highly dynamic and

variable in terms of their three-dimensional structure, mechanical properties and biochemical

microenvironment (Huh, Hamilton, & Ingber, 2011). The utilization of the third dimension,

provides the capability to layer cells onto cells. The architecture of blood vessels is a layer-by-

layer approach of endothelial cells. The development and implementation of this specific layering

of cells provides vasculature within microfluidic chips. Building on this design, the implementation

of other cells for the desired target function have led to the development of heterogeneous

microfluidic platforms; blood vessels (M. Shin et al., 2004; Song et al., 2005), muscles (Lam,

Huang, Birla, & Takayama, 2009), bones (K. Jang, Sato, Igawa, Chung, & Kitamori, 2008), airways

(Huh et al., 2007), liver (Huh et al., 2010; Y. Kang, Sodunke, Cirillo, Bouchard, & Noh, 2013;

Khetani & Bhatia, 2008; P. J. Lee, Hung, & Lee, 2007; Powers et al., 2002), brain (Park, Vahidi,

Taylor, Rhee, & Jeon, 2006), intestine (Mahler, Esch, Glahn, & Shuler, 2009) (Kimura, Yamamoto,

Sakai, Sakai, & Fujii, 2008), cornea (Figure 1-4)(N.-T. Nguyen et al., 2013; Puleo, Ambrose,

Takezawa, Elisseeff, & Wang, 2009), and kidney (K.-J. Jang & Suh, 2010). By applying micro-

technology, it is possible to unite simplicity and realism in one in vitro tissue model. The first

generation of organs-on-chips have demonstrated that there is a great potential to change the

18

landscape of in vitro testing for fundamental biology, drug development and toxicology (van der

Meer & van den Berg, 2012).

Figure 1-4. Adopted from Nguyen et al, this schematic illustrates a drug delivery microfluidic device implanted onto an eye (N.-T. Nguyen et al., 2013).

1.3 Advantages of Cell-laden Microfluidic Chips and Maskless Fabrication

Microfluidics, the science and engineering of fluid flow in micro-scale, is the enabling

underlying concept for microfluidic technologies (N.-T. Nguyen et al., 2013). Fluid dynamics on

a micro-scale level have paved the way for many biological benefits. The micro-architecture of

microfluidic chips have enabled laminar fluid flow through the designed platform. The

development of three-dimensional tissue scaffold with complete vasculature have proven to be a

difficult task. Without perfusion throughout a tissue scaffold, it is nearly impossible to deliver

nutrients to cells deep inside the scaffold (Linda G Griffith & Naughton, 2002). Since

manufacturing limitations have limited the design of implementing a method of perfusion in three-

dimensional scaffolds, researchers have turned to the manipulation of geometric constraints and

fluid mechanics to deliver a method for perfusion. In fluid mechanics, Reynolds number (Re) is

19

used to predict the flow pattern in any medium. Since Reynolds number is defined as the ratio of

the inertial forces to viscous forces, in micro-scale ‘pipes’; Reynolds number claims that the fluid

flow should be laminar. Laminar fluid flow is beneficial in biological constructs, in that it does not

allow for any mixing of nutrients and does not damage any particulates during transport.

Additionally, laminar fluid flow allows for the development of simple predictive mass transport

model which are developed to produce an array that enables perfusion throughout the desired

construct. Microfluidics have the potential to produce what is known as capillary fluid flow.

Capillary fluid flow does not require a driver to perfuse fluids throughout the construct. Many

microfluidic chips are designed to utilize the capillary fluid flow to enable perfusion (Whitesides,

2006). This method eliminates the need for a driver and the developed ‘patch’ which will self-

perfuse once implanted.

A close examination of any given tissue or organ in the human body will reveal a complex

mix of various cell types coupled together to accomplished the sole function of the targeted tissue

or organ. It is the integration and migration of these cells that are partially responsible for the

survivability of the tissue as cells proliferate and die throughout the tissue. Cell migration and

integration play a crucial role in various biological processes, including; embryogenesis, wound

healing, immune response, and tissue development (Nie et al., 2007). Understanding cell

integration and migration may lead to understanding and developing a tissue model that allows for

closer mimicry. Microfluidic chips allows for a simple investigation of how cells integrate and

migrate under micro-environment. This micro-environment can be tailored to mimic that of the

targeted tissue. Since microfluidic chips provide laminar fluid flow perfusion, layering and

mimicking the layout of the cell pattern in vitro as it is in vitro provides adequate data that allows

for the development of complex heterogeneous microfluidic chips that models a target function of

a tissue or organ. These models allow for the investigation of an invasive disease cell, such as

20

cancer onto a healthy tissue and the development of biological devices such as ‘patches’(N.-T.

Nguyen et al., 2013).

Microfluidic fabrication has been developed with two sets of materials: silicon or glass and

polymers. Silicon and glass have well-controlled mechanical and chemical properties but they also

have high manufacturing costs and high processing complexity, particularly for disposable devices.

By contrast, polymers can easily be fabricated via soft lithography or hot embossing, where a single

mold can serve as a template for many devices (Neuži, Giselbrecht, Länge, Huang, & Manz, 2012).

There are various methods of fabricating microfluidic chips, all of which utilizes some form of

lithographic process and are limited by material selection. The development of cell-laden

microfluidic chips cannot utilize any processes or materials that are not biological compatible.

These constraints have significantly hinder the fabrication of cell-laden microfluidic chips, let alone

cell-laden microfluidic chips. Additive manufacturing approaches have provided techniques to

fabricate cell-laden microfluidic chips by eliminating the need for long fabrication processes, the

use of a photo-mask, and the use of toxic chemicals, while allowing for spatially controlled

heterogeneous deposition of cells/biologics as the tissue array is being fabricated. This non-

conventional fabrication approach make investigations more economic; requiring shorter

fabrication time, less material to produce a construct, less cells due to its capability to deposit/print

cells directly into the micro-channels during the fabrication process and above all it will develop

microfluidics that allows for consistency in experimental analysis due to limited interactions with

end users (Hsiao et al., 2009; P. J. Lee, Gaige, Ghorashian, & Hung, 2007; Ong et al., 2008;

Tannock, Lee, Tunggal, Cowan, & Egorin, 2002; Toh et al., 2009; Toh, Ng, Khong, Samper, &

Yu, 2005; Tourovskaia, Figueroa-Masot, & Folch, 2005; A. P. Wong, Perez-Castillejos,

Christopher Love, & Whitesides, 2008).

21

Conventional manufacturing techniques utilized to fabricate microfluidic chips have used

global exposure where the entire sample is treated all at once with one uniform energy distribution.

This method does not allow for localized changes on the fabricated sample. Additionally, many

photo-sensitive material used for the development of microfluidic chips are not biologically

compatible. Some material allow for special surface treatment to allow for bio-compatibility. In

this, a global surface treatment is used. Global surface treatment does not allow for specialize

functionalization to target specific cell attachment at a targeted area(s). Coupling localized

treatment with additive manufacturing approaches allows for the potential to precisely fabricate

each area(s) of a microfluidic chip which models a closer mimicry of the target organ. Additionally,

localized treatment allows for the fabrication of cell-laden microfluidic chips. Since harmful

ultraviolet (UV) is localized, manufacturing techniques can be manipulated to enable a chip

fabrication where cells are precisely deposited at a targeted area(s) within the micro-channels of

the chip. Also, localized exposure and surface treatment eliminates the need for photo-mask;

making the fabrication process more economical. A fully automated fabrication process limits

human contact and significantly reduces human-errors.

1.4 Research Objectives and Approach

The objective of this research is to investigate the integration of maskless fabrication, direct

cell deposition, and surface modification techniques to engineer cell-laden microfluidics. This

thesis presents; advances in additive manufacturing techniques, the utilization of plasma chemistry

to enhance surface functionalization, and manipulation of photo-polymerization to investigate new

approaches to assemble cell-laden microfluidics. Specifically, this thesis scrutinizes the following

activities:

22

1) A study of SU-8’s potential to serve as a biologically compatible material for the

development of microfluidic chips with enhanced cell attachment and proliferation.

2) An inspection of utilizing a digital mirroring system with a multi-nozzle biologics

deposition system to assemble cell-laden microfluidics.

3) The exploration of a freeform micro-plasma system for the development of a three-

dimensional cell-laden microfluidic chip.

4) The development, implementation, and characterization of an additive fabrication

system which utilizes; a multi-nozzle biologics component for precise spatial printing

of cells, a micro-plasma head for localized surface functionalization, an ultra-violet

component for freeform exposure of photo-polymers, and a photo-polymer material

delivery component for direct deposition and fabrication of a three-dimensional micro-

architecture.

5) The development and characterization of a cell-laden microfluidic chip to investigate

drug metabolism and deliver chip that produces a microfluidic environment which

facilitates co-culture of cancerous cells.

1.5 Thesis Outline

This thesis is outlined as follows:

Chapter 2 presents a study of SU-8’s potential to serve as a biologically compatible

material for the development of microfluidic chips with enhanced cell attachment and proliferation.

The focus of this chapter is to enhance the chemical group functionality, surface charge,

hydrophilicity, hydrophobicity, and wettability of bare SU-8. Three surface treatments frequently

used in tissue engineering and regenerative medicine are investigated; 1) Plasma treatment, 2)

chemical reaction, and 3) deposition treatment. This chapter investigates these surface treatments

23

by characterizing their corresponding water contact angle, topology condition, chemical

distributions, and ability for cells to attach and proliferate. These investigations are presented as

evidence to effectively select the treatment that will be most beneficial for enhancing the biological

properties of SU-8.

Chapter 3 presents an inspection of utilizing a digital mirroring system with a multi-nozzle

biologics deposition system to assemble cell-laden microfluidics. This chapter describes the

process of utilizing a digital mirroring system to fabricate a cell-laden microfluidic chip. It also

demonstrates the capabilities of using a cell printing system to deposit cells into the micro-channels

of a chip. The assembled microfluidic chip is then characterized to illustrate the benefits of direct

cell deposition into microfluidic chips, cell morphology within the micro-channels of the chip, and

the chip’s structural integrity and cytotoxicity.

Chapter 4 presents the exploration of a freeform micro-plasma system for the development

of a three-dimensional cell-laden microfluidic chips. With the incorporation of a freeform micro-

plasma system, this chapter demonstrates the capabilities of fabricating a three-dimensional

interconnected microfluidic chip. Computational analysis is supplied as evidence to prove that the

fluid dynamics within the fabricated chip is still laminar and interconnected. This chapter also

presents the interconnected cell-laden microfluidic chip’s capabilities to serve as a platform to

investigate drug metabolism.

Chapter 5 presents the development, implementation, and characterization of an additive

fabrication system which utilizes; a multi-nozzle biologics component for precise spatial printing

of cells, a micro-plasma head for localized surface functionalization, an ultra-violet component for

freeform exposure of photo-polymers, and a photo-polymer material delivery component for direct

deposition and fabrication three-dimensional micro-architecture. Each component of this additive

fabrication system is characterize to enable its end-user to predictively model the fabrication

24

process to develop the desired chip. This chapter also presents the development and

characterization processes of a cell-laden microfluidic chip that provides a microfluidic

environment which facilitates co-culture of cells.

In closing, Chapter 6 presents the conclusions and recommendations for future work of this

research.

25

CHAPTER 2: SURFACE MODIFICATION OF SU-8 FOR ENHANCED CELL ATTACHMENT AND PROLIFERATION

2.1. An Inspection of Surface Modification

Biomaterials for life science applications are classified in two categories: 1) Natural

Biopolymer and 2) Synthetic Biopolymers. Natural biopolymers are polymers that occur in nature

and can be exacted and synthesized. Examples of natural biopolymers include, but are not limited

to, gelatin, alginate, and collagen. On the other hand, synthetic biopolymers are polymers that are

biocompatible (low levels of cytotoxicity and increased hydrophilicity) and a made by scientists

and engineers. Examples of synthetic biopolymers include, but are not limited to; polylactic acid

(PLA), hydroxyapatite (HA), and polystyrene. An important aspect of tissue construct fabrication

is the selection of the ideal biomaterial (Hutmacher, Schantz, Lam, Tan, & Lim, 2007). Apart from

the architectural design, the material used for the tissue construct fabrication has a bearing effect

of the mechanical, chemical, and biological properties (J. Y. Wong, Leach, & Brown, 2004). In

terms of the mechanical properties, the Young’s modulus of the scaffold is affected by the material.

For some tissue constructs, the tensile and compressive strengths are essential for that construct to

be successful (Discher, Janmey, & Wang, 2005; Yeung et al., 2005). For example, if a construct

is made from a low compressive strength material and is place in a highly compressive

environment, the construct will collapse and fail. Studies have also shown that the mechanical

properties have some effect on the cell’s abilities to attach and proliferate. The chemical and

biological properties influence the cell’s ability to attach, proliferate and differentiate. A

biomaterial may be biocompatible at it raw/stock (unprocessed) form, however, once processed its

properties can change (depending on the manufacturing techniques). Some manufacturing

processes can change the material’s toxicity level, making it difficult for cells to attach and

proliferate. In additional to toxicity, surface properties can be modified. Surface modification can

26

lead to a smooth or rough surface condition or it can change the degree of hydrophilicity or

hydrophobicity of the biomaterial.

In the late 1900s, the field of tissue engineering and regenerative medicine was established

to address the limitations of tissue grafting and tissue repair (Audet, 2004; Bonadio, 2000; Caplan,

Reuben, & Haynesworth, 1998; Cutroneo, 2003; Hollister, 2005; Langer & Vacanti, 1993;

Torquato, 2001). A major challenge of this is the ability to find materials and techniques that

promote cell attachment, proliferation, differentiation, and have specific architecture that enables

the development of an extracellular matrix (ECM) (F. Berthiaume, P. V. Moghe, M. Toner, & M.

L. Yarmush, 1996; L. G. Griffith, 2002; D. Han & Gouma, 2006; B. S. Kim & Mooney, 1998; Z.

W. Ma, Gao, Gong, & Shen, 2005). The ECM plays a critical role in the initial development of a

tissue array as it serves as the platform for which the architecture, topology, chemical composition,

and functional groups provide the proper environment for cells to attach and proliferate into

functional tissue construct (Cancedda, Dozin, Giannoni, & Quarto, 2003; Hollister, Levy, Chu,

Halloran, & Feinberg, 2000; W. J. Li et al., 2005; Ochi, Uchio, Tobita, & Kuriwaka, 2001; Oyane

et al., 2005; Tuan, Boland, & Tuli, 2003; Tuli, Li, & Tuan, 2003). Given that the success of a tissue

construct requires an ECM that mimics that target organ, the appropriate cell source, and optimal

signals for cell functioning, the design and fabrication of the ideal tissue array is very complex and

not yet fully understood (F. Yang, Wolke, & Jansen, 2008). Materials have been developed with

special properties that have attractive qualities to aid in the development of an array that closely

mimics the in vivo conditions of the target organ (W. J. Li et al., 2005).

The architecture, topology, and surface chemistry play an important role in the

development of a functioning tissue array. It is a challenge to fabricate constructs that mimic the

ECM’s structures with defined shapes and complex architecture(D. Han & Gouma, 2006). In

response, a close estimate can be made where, a biomaterial is chosen that can be fabricated to

27

provide the appropriate cues (Frame, Fincham, Carragher, & Wyke, 2002; Iivanainen, Kahari,

Heino, & Elenius, 2003; Larsen, Tremblay, & Yamada, 2003; M. Y. Li et al., 2005; L. L. Nguyen

& D'Amore, 2001; Nishimura et al., 2003; Suzuki et al., 2003). The topology of the tissue construct

has been found to affect cell morphology, differentiation, functionality, and physiological

responsiveness (F. Berthiaume, P.V. Moghe, M. Toner, & M.L. Yarmush, 1996; D. Han & Gouma,

2006). Besides the topology, the surface chemistry is also crucial as it provides the direct contact

with the surrounding cells and tissues (F. Yang et al., 2008). Effective surface modifications such

as plasma-ion beam treatment, electric discharge, surface grafting, chemical reaction, vapor

deposition of metals, and flame treatment (Williams, Martin, Horowitz, & Peoples, 1999; X. S.

Yang, Zhao, & Chen, 2002) change the chemical group functionality, surface charge,

hydrophilicity, hydrophobicity, and wettability (Liu, Jen, & Chung, 1999; Sacristan, Reinecke, &

Mijangos, 2000; X. S. Yang et al., 2002). A tissue construct with the appropriate mechanical,

chemical, and biological cues holds tremendous promise (Hollister et al., 2000).

SU-8 is a simple epoxy-based negative photoresist that was originally developed for

photolithographic manufacturing processes in the semiconductor industry. SU-8 has been

primarily used for structural elements and microfluidic components in MEMS. Literatures have

shown that SU-8 has mostly been used as micro molds or to fabricate freestanding and mechanical

structures (Despont et al., 1997; Genolet et al., 1999; Nordström, Marie, Calleja, & Boisen, 2004).

SU-8’s chemical, thermal resistance, high aspect ratio, and ability to produce a wide range of

patterned thicknesses makes this material a potential biomaterial for the development of a variety

of biological applications which include, but are not limited to, tissue engineering, drug delivery,

cell-based screening and sensing (Chang-Yen, Eich, & Gale, 2005; Del Campo & Greiner, 2007;

M. Evans, Sewter, & Hill, 2003; Jenke, Schreiter, Kim, Vogel, & Brugger, 2007; K. Lee et al.,

1995; Mata, Fleischman, & Roy, 2005; Stroock & Whitesides, 2002; Tao, Popat, Norman, & Desai,

28

2008). Since the native SU-8 surface is highly hydrophobic and has a low surface energy, alone, it

does not provide the appropriate cues necessary to support cell attachment, proliferation, and

differentiation (Calleja et al., 2005; Merz & Fromherz, 2005; Ribeiro, Minas, Turmezei,

Wolffenbuttel, & Correia, 2005; Walther et al., 2007). However, recent developments within the

tissue engineering and regenerative medicine field, allow for the modification of SU-8’s chemical

group functionality, surface charge, hydrophilicity, hydrophobicity, and wettability (Nordström et

al., 2004)

The focus of this chapter is to enhance the chemical group functionality, surface charge,

hydrophilicity, hydrophobicity, and wettability and develop a new technique (manufacturing and

surface modification) that allows for the development of cell-laden microfluidic chips. Three

surface treatments frequently used in tissue engineering and regenerative are investigated on SU-

8; 1) Plasma treatment, 2) chemical reaction, and 3) deposition treatment. O2 plasma treatment of

polymer surfaces yields completely wet-able surfaces with water contact angles of less than 5° and

modifies the surface to include oxygen-containing functional groups (Oyane et al., 2005; Walther

et al., 2007). Many cell culture protocols require the deposition of gelatin prior to seeding cells.

Gelatin is said to provide the appropriate cues necessary for cells to attach and proliferate; similar

properties are expected with the use of SU-8 (Marin, Kaplanski, Gres, Farnarier, & Bongrand,

2001; Paguirigan & Beebe, 2006). The third surface treatment chemically changes the surface

properties of SU-8. Literature surveys have shown that SU-8 can be chemically enhanced for

biological benefits (Nordström et al., 2004; Walther et al., 2007) This chapter will study these

three surface treatments and determine which surface treatment is the most beneficial for enhancing

the biological properties of SU-8 by characterizing each treatment’s wettability, topological

conditions, chemical composition, and biological potential to attach and proliferate cells

effectively.

29

2.2. The Development of a Bare SU-8 Substrate

All samples characterized in this chapter were fabricated from an in-house digital

microfabrication system. This microfabrication system is a Digital Light Processing (DLP) unit

that projects images of ‘.jpeg’, ‘.bitmap’, or ‘.gif’ formats. The micro-mirrors have the option to

switch between masks within a matter of micro-seconds while offering high resolutions

performance in Spatial Light Modulation (SLM). The main component is the digital micro-mirror

device (DMD), an optical semiconductor module that allows for the digital manipulation and

projection of UV light. The digital mirrors are mounted directly above the platform and are angled

towards the UV light source, which emits UV light, adjustable in terms of intensity and exposure

time. During the projection phase, the digital mirrors would either be ‘on’ or ‘off’ depending on

the pattern being projected. Mirrors that are turned ‘on’ would absorb the UV light and project it

downwards onto the substrate, while mirrors that are turned ‘off’ would reflect the UV light in the

opposite direction (T. Nederman, H. Acker, & J. Carlsson, 1983). Figure 2-1 shows a schematic

of the digital micro-mirroring microfabrication system.

30

Figure 2-1. A schematic of the digital micro-mirroring microfabrication system.

A multi-nozzle biologics printer was utilized for the deposition of cells into the micro-

channels of the fabricated chips. The biologics printer operates with the cell-friendly conditions of

room temperature and low pressure conditions. This system consists of three motion arms for three-

dimensional spatial control and a material deposition system which houses up to four biological

materials at once. The deposition system utilizes a micro-valve nozzle system that can deposit

numerous solutions with a wide range of material and biological properties. The computer

controlled multi-nozzle biologics deposition system eliminates human errors and provides its users

with precision control during fabrication procedures (R. Chang, Sun, W.,, 2009; W. Sun, Darling,

Starly, & Nam, 2004a; W. Sun & Lal, 2002a). Cell printing is considered to be an effective tool in

the field of tissue engineering to assemble biologics. Figure 2-2 illustrates a schematic of the multi-

nozzle biologics deposition system.

31

Figure 2-2. Schematic of the multi-nozzle biologics deposition system (R. Chang, Sun, W.,, 2009).

Polydimethylsiloxane (PDMS) (Dow Corning, Michigan, USA) is used as the base of the

chip while SU-8 2100 (MicroChem Corp., Newton, MA, USA) is used to fabricate the micro-

channels of the chip. Fabricated entirely from PDMS, the enclosure of the chip is developed to

house the micro-channels. The enclosure comprises of a platform (bottom) and a lid (top). The inlet

and outlet ports are nylon based luer-lock port (McMaster-Carr, Robbinsville, NJ, USA). PDMS

is mixed at 1:15 ratio, de-gassed and cured in an aluminum mold at 130°C for 10 minutes. The

cured PDMS is cooled and removed from the aluminum mold. This process is repeated for the lid

where the luer-lock ports are placed into position prior to being cured on the hot plate. Figure 2-3A

illustrates a model of the PDMS enclosure.

32

Figure 2-3. (A) Model of the PDMS enclosure, (B) micro-channel fabricated within the bottom enclosure of the chip.

The fabrication of the micro-channels started by pouring and leveling SU-8 within the

PDMS slot (bottom of the enclosure). The bottom enclosure with the SU-8 is soft-baked at 65°C

for 20 minutes, then at 90°C for 220 minutes for stability. Immediately after soft-baking, it is cooled

for 30 minutes then exposed at the recommended exposure time based on the amount of energy

required for crosslinking (provided by the manufacturer). The exposure time with the use of the

digital mirrors is 10.75 minutes, a total exposure of 557 mJ. The exposed sample was then hard

baked at 65°C for 15 minutes, then at 90°C for 30 minutes for structural integrity. Prior to

development with the SU-8 Developer (MicroChem Corp., Newton, MA, USA), samples are

cooled for another 30 minutes. During the development process, all unwanted SU-8 is washed

away. The total development time per sample is 8-15 minutes. After development, samples are

removed and rinse with deionized (DI) water to remove any excessive materials within the

channels. Figure 2-3B is an image of the actual micro-channel fabricated within the bottom

enclosure of the chip. Chips presented in this chapter have a continuous channel that is 300 µm

wide and 500 µm deep.

Prior to cell deposition within the micro-channels, all samples were surface treated and

sterilized. The three surface treatments investigated are; 1) Plasma treatment, 2) chemical reaction,

33

and 3) deposition treatment. The plasma treated chips were treated with a Harrick Plasma Treater

(Harrick Plasma, Ithaca, NY, USA). Each sample was vacuumed to a pressure of 100 mTorr to 1

Torr and plasma treated at high RF (18 W) power settings for 120 seconds. The second surface

treatment involves sulfuric acid. After development, chips were submerged in 99% sulfuric acid

at 80℃ (Sigma-Aldrich, USA) for 10 seconds. Chips were then removed and rinsed with DI water

to remove all unwanted materials and chemicals. The final surface treatment was the deposition of

2% (w/v) gelatin (bovine) (Sigma-Aldrich, USA). 0.5 mL of gelatin was uniformly spread onto

the micro-channels. Chips where then placed on a hot plate at 80℃ for dehydration. After

treatment, the multi-nozzle biologics printer deposits cells within the channels. Once the cells were

deposited within the micro-channels of the chip, the lid of the enclosure is placed on top of platform

(with micro-channel) to produce the cell-laden microfluidic chip. Prior to closing the chip, the lid

of the enclosure was plasma treatment to create a seal between enclosures. The cell-laden chip was

immediately placed in the incubator for 14 days in which biological characterizations are conducted

at various time points. Figure 2-4 shows and image of the multi-nozzle biologics deposition

system printing cells within the channels of the chip.

Figure 2-4. The multi-nozzle biologics deposition system printing cells within the channels of the chip.

34

Biological investigations used MDA-MB-231 cell line obtained from ATCC. Unless listed

otherwise, all cell culture supplements were obtained from ATCC. The MDA-MB-231 cell line

was seeded onto 75 cm2 vented flasks and incubated at 37 °C with 100% air. Six hours after the

cells were seeded, the culture medium was changed to remove any dead cells within the tissue

culture flask; culture medium was also changed every 2-3 days until flasks were confluent.

Confluent flasks are then harvested and counted using hemocytometer. Cells were then re-

suspended to a cell density of 1x106 cells/mL and then loaded into the cell printer where it’s printed

into the micro-channels. After the printing process, the chip was placed into the incubator with a

fluid line connected to the inlet and outlet of the chips. Culture medium is pumped through the

chips with the use of a syringe pump at a flow rate of 30 µL/hr. All biological investigation data

in this chapter are expressed as the mean ± standard deviation for sample size of 3 (n=3).

2.3. Water Contact Angle Investigations

Extensive characterization was conducted on four microfluidic samples: 1) untreated (bare

SU-8, no surface treatment), 2) 2% gelatin (deposition surface treatment), 3) air plasma (plasma-

ion surface treatment), and 4) 99% sulfuric acid (chemical surface treatment. Since cells prefer

hydrophilic substrates to attach onto and proliferate, a water contact angle (WCA) study was

conducted to investigate the wettability and hydrophilic/hydrophobic nature of each sample’s

surface (Grinnell & Feld, 1982). The contact angle is defined as the angle between the substrate

support surface and the tangent line at the point of contact of the liquid droplet with the substrate.

A drop of 2 µL Di-water was added to the center of each sample and measured by an in-house

goniometer that utilized a Basler A601f camera (Basler Vision Technologies) and a Fiber-Lite MI-

150 light source (Dolan-Jenner industries). Side-view images were taken of each sample to analyze

35

the WCA (shown in Figure 2-5). Drop shape analysis plug-in on ImageJ software was used to

characterize the WCA of each sample. The drop shape analysis plug-in quantifies the contact

angle based on the fitting of the Young-Laplace equation to the image data.

Figure 2-5. Side-view images showing water droplet illustrating the water contact angle on: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces.

Table 2-1 presents the results of the WCA investigation. According to the data, the bare

SU-8 is a hydrophobic material with an average WCA of 103.84° (Figure 2-5A). All surface

treatment changed the hydrophobic SU-8 to hydrophilic. The greatest change in hydrophilicity was

seen with the air plasma treatment where the average WCA was found to be 15.76° (Figure 2-5C).

Even though the sulfuric acid treatment changed the WCA of the bare SU-8 from a hydrophobic

surface to a hydrophilic surface, the WCA of 81.66° is a low hydrophilic surface (Figure 2-5D).

Since many medical applications have used some form of gelatin for its hydrophilic properties

(Takahashi, Miyoshi, & Boki, 1993), the author expected the gelatin treated samples shows a great

change in hydrophilicity with an average WCA of 45.08° (Figure 2-5B).

36

Table 2-1. Water Contact Angle Measurement.

Surface Treatment Water Contact Angle

Untreated 103.84° ±3.27°

Gelatin (2%) 45.08° ±3.04°

Plasma (Air) 15.76° ±1.52°

Sulfuric Acid (99%) 81.66° ±4.24°

2.4. Topological Analysis

Quantitative and qualitative topological characterizations were performed on each sample

using an optical profiler (Zygo NewViewTM 6000). The Zygo uses scanning white light

interferometry to image and measure the micro structure and topography of surfaces in three

dimensions. The optical microscope lateral resolution measures from 0.45 µm to 11.8 µm with a

data scan rate of up to 85 µm/sec. The height resolution is 0.1 nm. Table 2-2 shows the quantitative

results of the surface roughness of each sample where the peak-to-valley (P-V) gives a general idea

of the topological conditions featured on each surface. Additionally, the arithmetic average of the

roughness profile (Ra) and Root Mean Square (RMS) is tabulated in Table 2-2. Based on the P-V,

the plasma treated samples have the smoothest surfaces (P-V of 1.309 µm) while the sulfuric acid

treated surfaces had the highest P-V (P-V of 16.049) measurement. A 5.010 µm P-V measurement

of the untreated samples illustrates that the plasma treatment reduces the surface spikes. On the

other hand, it seems as if the sulfuric acid treatment etches the surfaces creating deeper valleys.

According to the P-V values for the gelatin treated samples presented in Table 2-2 and based on

the fact that the gelatin surface treatment is a deposition-based treatment, the valleys of the surfaces

are being filled to reduce the roughness. The data present in Table 2-2 cannot provide a clear

37

understanding of the topological conditions of each sample, hence, a three-dimensional surface

profile and a line profile were generated to future understand the exact surface conditions developed

by the respective surface treatment.

Table 2-2. Quantitative Analysis of Each Surface Treatment.

Surface Treatment P-V [µm] RMS [µm] Ra [µm]

Untreated 5.010 0.262 0.177

Gelatin (2%) 2.008 0.064 0.050

Plasma (Air) 1.309 0.034 0.027

Sulfuric Acid (99%) 16.049 0.667 0.359

The three-dimensional surface profile (Figure 2-6) and the line surface profile (Figure 2-7)

presents a vivid understanding of the topological conditions of each sample. The Zygo is optimized

to analyze an area of 500 µm x 700 µm from each sample. As seen on the three-dimensional profile,

each surface has a unique profile. According to the WCA results, the untreated surface is the most

hydrophobic sample. This is confirmed by Figure 2-6A which shows an extremely spikey profile.

In comparison, Figure 2-6C is the plasma treated sample (the most hydrophilic surface) whose

surface seems to be flat. The quantitative results presented in Table 2-2 suggest that the sulfuric

acid treated sample is very rough. After reviewing the three-dimensional surface profile, there are

isolated spikes and flat surfaces on the sulfuric acid treated samples (Figure 2-6D). This suggests

that the sulfuric acid reduces the spikey topology. However, with the varying profile of spikes and

flat surfaces, this suggests that a quick treatment of 10 seconds and no agitation may not have been

sufficient to produce a more hydrophilic profile. On the other hand, a longer treatment of sulfuric

acid will cause unwanted deformation to the micro-channels.

38

Figure 2-6. Three-dimensional profile of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces.

Figure 2-7 presents the line profile of each of the studied samples. On the line profile plot,

there is a dashed-line that does horizontally across the plot. This dashed-lined presents the base

(zero level) of the substrate. This base-line is tabulated at the beginning of each test by the

characterization instrument, this process is referred to as ‘homing and leveling’. Since the gelatin

treatment is a deposition treatment, the line profile (Figure 2-7B) coupled with the three-

dimensional surface (Figure 2-6B) profile will provide a clear understanding of the topological

conditions created by this treatment. According to the line profile presented in Figure 2-7, it is

confirmed that this treatment filled valleys. The combination of the chemical properties of gelatin

and the topological conditions produced by the deposition treatment creates a hydrophilic surface

situation for microfluidic chips. A closer look at the line profiles shows that sulfuric acid treatment

etches into the SU-8 (Figure 2-7D) while the other treatments does not (Figure 2-7A-C).

39

Figure 2-7. Line profile of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces.

2.5. X-Ray Photoelectron Spectroscopy (XPS) Analysis

To correlate the change in surface morphology with the local chemical composition, survey

and carbon 1s spectra of XPS were obtained from four samples under previously mentioned surface

different treatment methods. The results of the XPS survey spectra of the untreated surface is

presented in Figure 2-8, the gelatin treated surface is presented in Figure 2-9, the plasma treated

surface is presented in Figure 2-10, and the sulfuric acid treated surface is presented in Figure 2-11.

Figure 2-12 thru Figure 2-15 presents the results of the detailed XPS spectra of carbon 1s for

untreated (Figure 2-12), gelatin treated (Figure 2-13), plasma treated (Figure 2-14), and sulfuric

acid treated surfaces (Figure 2-15). The set of experiment and analysis was performed using a

VersaProbe II Scanning XPS Microprobe (Physical Electronics, Inc.) and its associated proprietary

software program MultiPak. All XPS spectra were obtained using a monochromatic Al Kα X-ray

source (1486.6 eV and 97.1 W). Table 2-3 presents the atomic elemental composition of each

treated sample surface from XPS survey spectra. Carbon, oxygen, silicon and chlorine were

40

observed on the untreated sample, which could be resulted from the chemical nature of SU-8 with

PDMS, and also from the manufacturing process of micro-channels. After each treatment

respectively, silicon and chlorine on sample surface were mostly removed or coated by carbon and

oxygen functional groups which would promote cell attachment and proliferation.

Figure 2-8. XPS survey spectra of untreated surfaces.

41

Figure 2-9. XPS survey spectra of the gelatin treated surface.

Figure 2-10. XPS survey spectra of the plasma treated surface.

42

Figure 2-11. XPS survey spectra of the sulfuric acid treated surface.

Figure 2-12. Detailed XPS spectra of carbon 1s for the untreated surface.

43

Figure 2-13. Detailed XPS spectra of carbon 1s for the gelatin treated surface.

Figure 2-14. Detailed XPS spectra of carbon 1s for the plasma treated surface.

44

Figure 2-15. Detailed XPS spectra of carbon 1s for the sulfuric acid treated surface.

Table 2-3. Atomic Elemental Composition of Each Treated Surface

Surface Treatment C (1s) [%] O (1s) [%] Si (2p) [%] Cl (2p) [%]

Untreated 45.9 32.1 19.9 2.1

Gelatin (2%) 68.0 32.0 0.0 0.0

Plasma (Air) 67.9 30.3 1.8 0.0

Sulfuric Acid (99%) 74.4 25.0 0.0 0.6

After curve fitting, the detailed XPS spectra of carbon 1s are mainly divided into three

peaks: carbon-carbon (C-C), ether carbon (C-O-C), and aldehyde or carboxyl carbon (C=O or O=C-

O). Full width at half maximum (FWHM) of C-C largely decreased in all three surface treated

samples compared to untreated one, while FWHM of C-O-C increased. The increase in C=O or

45

O=C-O peak may be resulted from the introduction and formation of carboxylic acids and aldehyde

groups on sample surface from treatment process.

2.6. Biological Investigations

Since the micro-channels are fabricated from a photo-material that is enhanced for

biocompatibility, it is important to characterize the cytotoxicity of the chip. The interactions of the

cells within the micro-channels are of interest. Healthy cells have the ability to attach to the

substrate and proliferate; this is an indicator that the substrate in which the cells are growing on is

not toxic. A fluorometric investigation was conducted which characterized the cell-cell interaction

and proliferation within the fabricated chips. Cell interactions within the channels play an

important role in the development of a cell-laden microfluidic chip (Koh, Yong, Chan, &

Ramakrishna, 2008). This biological characterization was performed with the use of AbD

SeroTEC’s Alamar Blue (Ab). Ab is a simple water soluble indicator dye designed to provide a

rapid and sensitive measure of cell proliferation and cytotoxicity. The cell-laden chips were washed

with 1x Phosphate buffered saline (PBS) by pumping the PBS through the chips with a syringe

pump at a flow rate of 30 µL/hr. 10% Ab was mixed with culture medium and was pumped through

the chips at 30 µL/hr until the chips were filled with the reagent. The chips were then disconnected

from the syringe pump and were placed in the incubator for 4 hours. After 4 hours, the resulting

reagent within the chips was removed from the chips and characterized with a micro-plate reader

(GENios, TECAN, North Carolina, USA) whose excitation and emission wavelengths were 535nm

and 590nm respectively.

As seen in Figure 2-16, there is some form of up-regulated cell proliferation present in each

sample. The plasma treated samples showed the most active cell proliferation trend-line throughout

the 14 day period (p<0.00001). Since bare SU-8 is not very hydrophilic and is not a good candidate

46

which can facilitate cell life, it is expected that the untreated samples did not show much in terms

of cell proliferation. Since gelatin is used in cell cultures, it was expected that this surface treatment

would have a high cell count at the end of the study, however, the data suggests otherwise

(p<0.00001) (Marin et al., 2001; Paguirigan & Beebe, 2006). At the end of the 14 day study, the

sulfuric acid treated chips had a higher cell count (in comparison to the untreated chips), which

suggests that the change of hydrophilicity and topological conditions does support cell growth

(p<0.00087).

Figure 2-16. Cell proliferation study of untreated, gelatin treated, plasma treated, and sulfuric acid

treated surfaces.

Cell morphology was evaluated using an FEI/Philips XL-30 Field Emission Environmental

Scanning Electron Microscope (SEM). The images obtained from the SEM were taken using a

beam intensity of 2kV and gaseous secondary electron detectors of 1.3 Torr. Chips were washed

twice with 1x PBS (pumped through the chips), then fixed with 4% glutaraldehyde (Sigma Aldrich,

USA) for 2 hours. After being fixed, each chip was subjected to dehydration by pumping series of

47

diluted ethanol (50%, 70%, 90%, 95%, and 100%) through each chip. After the dehydration

process, chips were sectioned, dried, and then refrigerated at 4°C for 24 hours. Since the ion beam

cannot penetrate the PDMS layer, sectioning the chip is the only way the view the cells within the

micro-channels. The method of sectioning does cause some deformation to the micro-channels.

Figure 2-17 consist of SEM images showing the cell morphology of: (A) untreated, (B)

gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces. The SEM image confirms

that the MDA-MB-231 cell line was able to attach to the substrate. The morphology of the cells

on the varying surface treatment differs. The cells on the plasma treatment surfaces are within

close proximity with each other and are well anchored to the substrate. The cells on the gelatin

treated substrate are flat in comparison to the other three surfaces. The cells on the sulfuric acid

treated and untreated surfaces seem isolated (cells are not in close proximity with each other);

however, the cells on the sulfuric acid were well anchored. The morphological study along with

the cell proliferation investigation confirms that the plasma treated samples are a better surface

enhancement for the development of cell-laden microfluidic chips. Although the sulfuric acid and

gelatin treatments demonstrated potentials of being a good biological enhancement of SU-8, their

potential was significantly lower than that of the plasma treated microfluidic chips.

48

Figure 2-17. Cell morphology of: (A) untreated, (B) gelatin treated, (C) plasma treated, and (D) sulfuric acid treated surfaces.

2.7. Interpretations

This chapter focuses on enhancing the chemical group functionality, surface charge,

hydrophilicity, hydrophobicity, and wettability of SU-8. SU-8’s chemical, thermal resistance, high

aspect ratio, and ability to produce a wide range of patterned thicknesses make it a potential

biomaterial for the development microfluidic chips for tissue engineering applications. The

architecture, topology, and surface chemistry all play an important role in the development of a

functioning tissue array. Microfabrication techniques possess the ability to control surface

microarchitecture, topography, and feature sizes necessary to develop tissue arrays that aim at

49

restoring, maintain, or improving tissue function. A tissue construct with the appropriate

mechanical, chemical, and biological cues holds tremendous promise.

Surface treatments investigated in this chapter are ones that are frequently used in tissue

engineering and regenerative medicine; 1) Plasma treatment, 2) chemical reaction, and 3)

deposition treatment. An untreated microfluidic chip was characterized for comparison. The WCA

investigation tabulated the hydrophilic/hydrophobic capabilities of each surface treatment. The

investigation concluded that the plasma treatment yields the most hydrophilic surfaces while the

untreated sample had the most hydrophobic surfaces. To further understand the topological

conditions created with the surface treatment, an optical profiler was used to quantify the surface

profile. The optical profiler confirms that the untreated surfaces were extremely rough while the

plasma treated and gelatin treated surfaces were smooth, the plasma treated surfaces being the

smoothest of the two. The sulfuric acid treated surfaces had isolated patches of spikes and flat

surface. It was then confirmed with the line profile that the sulfuric acid etches the SU-8.

A proliferation study was conducted on all samples to investigate the each treatment’s

ability to support cell life. The data of this study showed that cells preferred the plasma treated

surface. The untreated surface did not show a significant increase of cells over the 14 day study.

The gelatin treated surface had more cells than the sulfuric acid at the end of the 14 day study,

however the cell count of the plasma treated surfaces were significantly higher. SEM

characterization later confirmed that the surface treatment does have an impact on the cell

morphology. The investigations presented in this chapter demonstrated that the plasma, gelatin,

and sulfuric acid treatments have a potential to enhance SU-8’s surface for biological application.

Of course each treatment has their advantages and disadvantages (application dependent). The

plasma treated surface is suggested to be the better of the three treatments for biological

enhancement followed by gelatin and sulfuric acid treatments, respectively.

50

CHAPTER 3: UTILIZATION OF A DYNAMIC DIGITAL MICRO-MIRRORING SYSTEM WITH A MULTI-NOZZLE BIOLOGICS DEPOSITION SYSTEM TO

FABRICATE CELL-LADEN MICROFLUIDICS

3.1. Applications of a Digital Micro-mirroring System

There is an overwhelming need for substitutes to repair tissues and organs because of

disease, trauma, or congenital problems. In the US alone, as many as twenty million patients per

year suffer from various organs and tissue related maladies caused by burns, skin ulcers, diabetes,

and connective tissue defects which include bone and cartilage damage. More than eight million

surgical procedures are performed annually to treat these cases, over 70,000 people are on

transplant waiting lists, and an additional 100,000 patients die due to the lack of appropriate organs

(Almeida, Bártolo, & Ferreira, 2007; "The Organ and Transplantation Network," 2004; B. Starly,

Lau, Bradbury, & Sun, 2006). Scientists are working around the clock to develop pharmaceuticals

and tissue replacements that would allow humans to live longer lives. However, many of these

developments require tremendous investigation on its effects on humans. Quite often, the use of

animal and human models is limited by the feasibility of testing protocols, availability, and ethical

anxieties (Elliott & Yuan, 2011; Parnes, Sun, & Freeman, 1999). Micro-Electro-Mechanical

Systems (MEMS) technologies have been very attractive and demonstrate the potential for many

applications in the field of tissue engineering, regenerative medicine, and life sciences. These fields

bring together the multidisciplinary field of engineering and integrated sciences to fabricate tissue

models that aids the exploration, generation or regeneration of organic tissues and organs (Huang

et al., 2011; L. Shor, 2008; B. Starly, 2006). MEMS were first introduced on conventional

semiconductor materials, and since then, MEMS have been utilized in many other fields with great

success (Ho & Tai, 1998; Jo, Van Lerberghe, Motsegood, & Beebe, 2000; Spearing, 2000).

The digital micro-mirroring microfabrication (DMM) system gives scientists the

capabilities to develop models that can be utilized to characterize new pharmaceuticals, tissue

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replacements, and develop models to study fatal disease such as cancers and tumors (D. S. Cowan,

K. O. Hicks, & W. R. Wilson, 1996; Saadia B Hassan et al., 2001; X. Zhang et al., 2005a). This

biologically inspired microfabrication system has the potential to develop critical three-

dimensional models for the investigation of various tissue models and biological sensors. Three-

dimensional biological models are preferred for in vitro investigation since these models eliminate

the limitations of traditional mainstay two-dimensional models (J. J. Casciari et al., 1994; M. J.

Friedrich, 2003). The DMM has the capabilities to fabricate many advantageous devices. Amongst

them, microfluidics systems have the most tissue engineering, regenerative medicine, and life

sciences applications to develop in vitro tissue models.

Unlike conventional microfabrication techniques, the DMM eliminates the need for mask

by incorporating a dynamic maskless fabrication technique (Adeyemi, Barakat, & Darcie, 2009; Y.

Lu, Mapili, Suhali, Chen, & Roy, 2006; W. Shin et al., 2006; Xiang & Arnold, 2011). Since the

DMM system can develop models on a micro-scale level, this would make the fabrication of tissue

constructs and biological investigation more economic; requiring less reagents, cells, and allow for

consistency in experimental analysis to due limited interactions with the end user (Andersson &

van den Berg, 2004; Catros et al., 2012; Gauvin et al., 2012). The DMM system is specifically

designed for the developments of biologically inspired devices, which includes, but are not limited

to, biosensors, lindenmayer systems, and micro-organs. Figure 3-1shows and outline of the

application potential of the DMM. This fabrication system eliminates the limitations of

conventional photolithography and enables the end user with the capabilities to develop

advantageous models within minutes (Adeyemi et al., 2009; B. Starly, Sun, W.,, 2007).

This chapter focuses on the developing a microfluidic chip for cells to attach and proliferate

within its channels. Eventually, this model is enhanced, in which drugs is evaluated within the

microfluidic chip. As listed in Figure 3-1, chips developed in this chapter can be categorized as

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biosensors. The fabrication and cell seeding techniques are non-conventional approaches that

support enhanced cell attachment and growth within a microfluidic chip. A sinusoidal micro-

pattern is fabricated from SU-8 and is housed within a Polydimethylsiloxane (PDMS) enclosure.

The SU-8 channels are plasma treated to enhance the material’s bio-compatibility. Additionally,

the plasma treatment creates a PDMS-PDMS bond, this bond seals the chip. The sinusoidal pattern

demonstrates the DMM’s capabilities to create complex microfluidic architectures while

showcasing the cell printer’s potential to uniformly deposit cells within the microchannels. All

biological investigation data presented in this chapter are expressed as the mean ± standard

deviation for sample size of 3 (n=3).

Figure 3-1. Applications of the dynamic digital micro-mirroring microfabrication system

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3.2. Digital Micro-mirroring System

The DMM consists of three major components: 1) digital micro-mirroring projection

system, 2) photolithographic substrate alignment system, and 3) mask modeling system. The micro-

mirroring projection system is connected to a computer interface. The computer system activates

and deactivates the projection of the mask onto the substrate. The photolithographic substrate

alignment system consists of a digital microscopic device that allows for alignment of the

substrate’s features. The mask modeling system utilizes computer-aided design (CAD)

technologies to design mask for projection. Mask projected by the micro-mirrors must be in .jpeg,

.bitmap, or .gif formats. Figure 3-2 shows the control system diagram of the digital micro-mirroring

microfabrication system.

Figure 3-2. Structure of the digital micro-mirroring microfabrication system.

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The micro-mirroring projection system has the potential to switch between masks within a

matter of microseconds while offering high resolutions performance in Spatial Light Modulation

(SLM). With ultraviolet (UV) light, this system offers a flexible platform to design and develop

proof of concepts, tissue models, biosensors, micro-organs, and lindenmayer systems. An image

of the digital micro-mirroring microfabrication system is shown in Figure 3-3.

Figure 3-3. Digital micro-mirroring microfabrication system.

The main component of the digital micro-mirroring projection system is the digital micro-

mirror device (DMD): an optical semiconductor module that allows the digital manipulation and

projection of UV light. The DMD comprised of millions on micro-mirrors aligned in a rows and

columns setting. During the projection phase, mirrors of the DMD would either be on or off

depending on the pattern being projected. Mirrors of the DMD that are turned on would absorb the

UV light and project it downwards, while mirrors that are turned off would reflect the UV light in

the opposite direction (T. Nederman et al., 1983). Figure 3-4 shows the direction of the light

projected onto the micro-mirrors and their corresponding reflection. The DMD is mounted directly

above the alignment platform and is angled towards the UV light source. The DMD is completely

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adjustable in terms of elevation and angle. The UV light source emits an adjustable light in terms

of intensity and exposure time. The light from the UV lamp travels and uniformly distributes on

the micro-mirrors. Energy is lost during transmission of light from the UV source to the digital

micro-mirror and from the micro-mirror to the substrate. The transmission wavelength of the

system ranges from 370 nm to 410 nm (Figure 3-5A) while the maximum energy output of UV

Source is 15 W at 100% intensity (Figure 3-5B).

Figure 3-4. Illustration of light reflection on the digital mirrors.

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Figure 3-5. (A) The digital micro-mirroring transmission spectrum, (B) The Ultraviolet source relative intensity range.

The photolithographic substrate alignment system consists of two actuators (manual); one

that controls motion in the X direction (Cartesian coordinates) and the other that controls motion

in the Y direction (Cartesian coordinates), together these two actuators span the X-Y plane of the

alignment platform. Along with the X-Y actuators, there is a pair of fasteners that holds the

substrate in place. Since this a microfabrication device, any small movement can be catastrophic.

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To ensure that the substrate’s features are aligned with the projected mask (from the micro-mirror

projection system) a digital microscopic device is directly below the alignment platform (centered)

with a live feed to the computer system. This microscopic device features a fully adjustable

magnification ranging from 20X to 200X. Additionally, the microscopic device has the capability

to adjust its focal distance. Figure 3-6 shows the control system diagram of the photolithographic

substrate alignment system.

Figure 3-6. Structure of the photolithographic substrate alignment system.

3.3. Multi-nozzle Biologics Deposition System

The multi-nozzle biologics deposition system is inspired by rapid prototyping technology

and is built on CAD/CAM platform, which is integrated with solid freeform automation to assemble

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biologics in three-dimensional space. This system consists of three motion arms for three-

dimensional spatial control and a material deposition which houses up to biological materials at

once. The deposition system utilizes micro-valve nozzle systems that can deposit a wide range of

solutions with a wide range of material and biological properties. This printer is fully integrated

and computer controlled. The multi-nozzle biologics deposition system eliminates human errors

and provides its end users with precision control during fabrication procedures. This system

executes micron-scale spatial control to generate cell-laden constructs. The multi-nozzle biologics

deposition system is capable of depositing heterogeneous materials, cell types, and biological

factors in a controlled and reproducible manner (R. Chang, Sun, W.,, 2009; W. Sun et al., 2004a;

W. Sun & Lal, 2002a). Cell printing is considered to be an effective biofabrication tool to assemble

biologics. An image of the major components of the biologics deposition system is shown in Figure

3-7.

Figure 3-7. An image of the major components of the Multi-nozzle Biologics Deposition System.

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A pneumatic micro-valve nozzle head has been selected to operate as the printer’s head

after extensive investigation to evaluate its performance and feasibility to deposit biologics

solutions for life sciences tissue constructs. The valve of this nozzle opens and closes when air

pressure is applied. The air pressure is regulated by the computer system; hence the computer has

full control of the micro-nozzle. There are a maximum of 4 nozzles that can be operated at the

same time, making this system a multi-nozzle deposition system capable of printing several

biologics at once. Figure 3-8 is a cross-sectional schematic of the pneumatic micro-valve nozzle

for the multi-nozzle biologics deposition system.

Figure 3-8. Pneumatic micro-valve nozzle for the multi-nozzle biologics deposition system.

3.4. Microfluidic Chip Fabrication and Characterization Protocols

Enclosure and Internal Architecture. The microfluidic chips are fabricated from two

materials. Polydimethylsiloxane (PDMS) (Sigma-Aldrich, St. Louis, MO, USA), is used as the

enclosure of the chip while SU-8 2100 (MicroChem Corp., Newton, MA, USA) is used to fabricate

the micro-architecture of the chip. The enclosure of the chip is fabricated first. There are two parts

of the enclosure; the platform (bottom) and the lid (top). The entire enclosure is fabricated from

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PDMS. The input and output ports are nylon based luer-lock port (source: McMaster-Carr,

Robbinsville, NJ, USA). PDMS is mixed at 1:15 ratio, de-gassed and cured in an aluminum mold

at 130°C for 10 minutes. Cured PDMS is cooled and removed from the aluminum mold. This

process is repeated for the lid, the luer-lock ports are placed into position prior to being cured on

the hot plate.

The DMM utilizes a systematic approach to fabricate models using a photosensitive

polymer. The digital micro-mirroring microfabrication system projects an image of the desired

structure onto the photosensitive polymer. Once the polymer is exposed, it manipulates the

material’s chemical properties and mimics the projected pattern (Guijt & Breadmore, 2008). Chips

presented in this chapter are fabricated using the following protocol. SU-8 is poured and leveled

within the PDMS slot (bottom of the enclosure). It is then soft baked at 65°C for 20 minutes, then

at 90°C for 220 minutes for stability. Cooled for 30 minutes then exposed at recommend exposure

time (this is based on the amount of energy required for crosslinking): DMM exposure time is

10.75 minutes, 557 mJ. The exposed sample is then hard baked at 65°C for 15 minutes, then at

90°C for 30 minutes for structural integrity. Sample is cooled for another 30 minutes then

developed with SU-8 Developer (MicroChem Corp., Newton, MA, USA); during this stage, the

unwanted material is washed away. Development time ranges from 8-15 minutes. Once developed,

the sample is removed and rinse with DI water to remove any excessive materials within the

channel.

Sterilization and Cell Printing. Chips are first sterilized, then plasma treated. All samples

used for biological investigations are sterilized first by applying dry heat of 150 °C for 3 hours.

Since the SU-8 and cured PDMS is thermally insensitive, sterilizing the chips with dry heat is

beneficial. Dry heat sterilization prevents moisture from being trapped in the microchannels

(compared to autoclave sterilization process). Prior to cell deposition within the microchannels,

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all samples are plasma treated with a Harrick Plasma Treater (Harrick Plasma, Ithaca, NY, USA):

vacuumed to a pressure of 100 mTorr to 1 Torr and plasma treated at high RF (18 W) for 120

seconds. Plasma treatment is used to create a seal between enclosures and enhances cell attachment

and proliferation within the SU-8 channels.

After the plasma treatment, harvested cells are loaded in the reservoir of the multi-nozzle

biologics deposition system. Cells are then deposited into the microchannels of the chips with an

applied pressure of 5 psi and a motion velocity of 1 mm/s. Cells are deposited into the channels

through a 250 µm nozzle. Once the cells are printed into the channels, the lid of the enclosure was

placed over the platform (base). The plasma treatment prior to the printing creates a PDMS-PDMS

bond, sealing the chip.

Cell Cultures. 7F2 (mouse osteoblast) (American Type Culture Collection, (ATCC),

USA) and MDA-MB-231(human breast cancer) (ATCC, USA) cell lines have similar cell culture

protocols. Both cell lines were seeded onto 75cm2 vented flasks and incubated. Six hours after the

cells were seeded, the culture medium (cell depended) was changed to remove any dead cells in

the flask; culture medium was also changed every 2-3 days until flasks are confluent. Confluent

flasks were harvested and counted using hemocytometer. Cells were then centrifuged again in

which the cell pallet was suspended to a cell density of 1x106 cells/ml. Cells are then loaded into

the printer where it’s printed into the microchannels. After the printing process, the open chips

(chips without lids) will be placed in the incubator for further characterization. The closed chip

would be sealed (sealing is possible to plasma treatment) immediately after cell printing with a

PDMS lid and placed in the incubator. Once the cells are attached onto the substrate (optical

verification), the open chips are placed in a petri dish and are submerged in culture medium. A

syringe pump is used the supply culture medium to the closed chips at a rate of 30 µL/hr. Culture

medium was change every 2-3 days in static culture (cell culture and open chips), after every

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characterization point and continuously for the dynamic culture (closed chips). 7F2 cells were

incubated at 37 °C with 95% air and 5% CO2 while MDA-MB-231 cells were at 37 °C with 100%

air. Unless listed otherwise, all cell culture supplements were obtained from ATCC, Manassas,

VA, USA.

Cell Interaction, Cytotoxicity Analysis, and Structural Integrity. A fluorometric indicator

(Alamar Blue, Serotec) of cell metabolic activity was utilized to determine the cell proliferation

within the channels of both treated and untreated chips (closed and open) (Q. Hamid et al., 2011;

L. Shor et al., 2009; K. C. Yan, Nair, & Sun, 2010). The open cell laden chips were removed from

the petri dishes, washed twice with 1x Phosphate buffered saline (PBS), placed into a new dish

where 10% (v/v) Alamar blue was added and incubated for 4 hours. The closed chips were washed

with 1x PBS by pumping the PBS through the chips with a syringe pump at a flow rate of 30 µL/hr.

10% Alamar blue was mixed with culture medium and was pumped through the chips at 30 µL/hr

until the chips were filled with the reagent. The chips were then disconnected from the syringe

pump and were placed in the incubator for 4 hours. After 4 hours, the resulting reagent from both

open and closed chips were removed and characterized with a micro-plate reader (GENios,

TECAN, North Carolina, USA) whose excitation and emission wavelengths were set at 535nm and

590nm respectively.

MarkerGeneTM Live:Dead cytoxicity assay kit was used to provide qualitative data of cells

within the microchannels. Manufacture’s protocols were followed to create the working live:dead

solution from the propidium iodide (PI) solution and the carboxyfluorescein di-acetate (CFDA)

solution. The carboxyfluorescein dye is retained within live cells, producing a green fluorescence,

while cells with damaged membranes allow the entrance of PI, which undergoes a fluorescence

enhancement upon binding to nucleic acids promoting a red fluorescence in dead cells. Qualitative

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data was collected from the cell-laden microfluidic constructs on day 7 after cells were printed in

the channels.

An additional cytotoxicity analysis was conducted to confirm the cell’s cytoplasm and

nucleus are not damaged from the printing process. This investigation was conducted 24 hours

after cells were deposited within the microchannels of the chip. The chips were prepped for

confocal microscopy by first sectioning the lid. Chips were then washed 3 times with 1x PBS and

stained with Calcein-AM (Dojindo, Japan, 1 μmol/L) and Propidium Iodide(Sigma-Aldrich, USA,

2 μmol/L) and incubated at 37 ℃ for 15 minutes. Calcein-AM is retained within live cells,

producing a green fluorescence while cells with damaged membranes allow the entrance of

Propidium Iodide promoting a red fluorescence. Prior to observation under the Laser Scanning

Confocal Microscope（Zeiss 710 META, Germany) chips were washed with 1x PBS to remove

the reagents.

Finally, cell morphology was evaluated using a FEI/Philips XL-30 Field Emission

Environmental Scanning Electron Microscope (SEM). Images taken by SEM used a beam intensity

of 2 KV and gaseous secondary electron detectors of 1.3 Torr. Chips were sectioned with a sharp

razor to remove the lid of the chip. The sectioned cell-laden chips were submerged in 2%

glutaraldehyde (GTA) (Sigma-Aldrich, St. Louis, MO, USA) for 2 hours followed by a dehydration

process of submerging the GTA treated samples in 70%, 80%, 90%, 95%, and 100% ethanol for

10 minutes, respectively. Following the dehydration process, the chips were placed under the

culture hood for 1 hour to dry then refrigerated at 4°C overnight. Prior to SEM, samples were

coated with Platinum for enhanced visibility.

3.5. Cell Proliferation, Cytotoxicity Analysis, and Cell Morphology

The microchannel array fabricated for this investigation is a single layered open and closed

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chip with a continuous channel whose dimensions are 300µm wide and 500µm deep. Both chips

characterized the chip’s potential for cell attachment and proliferation. There are two sample types

for the preliminary investigation; 1) non-plasma treated microchannels and 2) plasma treated

microchannels. Literatures have stated that plasma treatment enhances the surface properties of

biomaterials, the preliminary investigation will confirm if this holds true for SU-8 microchannels.

The cell printer is used to deposit cells within the microchannels of both open and closed chips.

7F2 is seeded in the open chip and MDA-MB-231 is seeded in the closed chip. These two cell lines

will demonstrate the DMM capabilities to develop micro-chips that can support viable diseased

(MDA-MB-231) and non-disease (7F2) cells.

It is important that cells maintain interaction within the microchannels; without the cell–

cell interaction, cells cannot proliferate and differentiate into mature cells that are essential for

functional tissues. An investigation was conducted on open and closed chips to characterize the

active proliferation within the microchannels for 14 days. Associated with cell proliferation is the

cell’s ability to attach onto the substrate. The 7F2 and MDA-MB-231 cell lines are like most cell

lines in which they need to attach themselves onto the substrate to actively proliferate. As stated

previously plasma treating the SU-8 enhances its bio-compatibility. In addition to an open and

closed chip proliferation study, there is a plasma treated open and closed chip proliferation study.

The plasma treatment study will confirm the effectiveness of plasma treating the SU-8 micro-

architecture. The results from this study are shown in Figure 3-9.

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Figure 3-9. 14 day cell proliferation study of treated and untreated open and closed microfluidic chips.

According to the data presented in Figure 3-9, the untreated open and closed chips showed

no significant up-regulation of cell proliferation. The plasma treated open chips with the 7F2 cells

showed a linear progression of proliferation up to day 7. After day 7, the progression subdued. The

micro-environment coupled with the limitation of nutritional supply within the microchannels

(typical in static culture) is believed to be responsible for the drop in cell proliferation of the open

plasma treated chips(E.A. Botchwey, M.A. Dupree, S.R. Pollack, E.M. Levine, & C.T. Laurencin,

2003; Leong et al., 2003). The closed chip with the MDA-MB-231 cell line showed a continuous

cell proliferation throughout the 14 day study. From days 3 to 7 both open and closed plasma

treated chips displayed similar trend-lines. The fluid flow within the closed chip is believed to be

responsible for the continued cell proliferation within the closed chip after day 7. The continuous

supply of culture medium within the microchannels of the chip is credited for the prolong cell life.

In comparison to the 7F2 cells used with the open chips, the MDA-MB-231 in the closed chip had

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a low initial cell count. The fluid flow through the microchannels of the closed chips may be

responsible for the low cell count at the beginning of the proliferation investigation.

This study confirms that plasma treatment of the SU-8 microchannels does enhance the

biocompatibility and is needed to develop cell-laden microfluidics. Additionally, the active

proliferation of both cell lines proved that these microfluidic chips can sustain viable diseased and

non-diseased cells. A dynamic culture (medium is actively flowing through the microchannels)

sustains a longer up-regulation of cell proliferation by supplying nutrients throughout the chip.

This allows for longer biological investigation and an increased number of cells per unit volume.

Conclusively, this investigation showed that closed plasma treatment microfluidic chips are a good

platform for cells. Since the closed plasma treated chips are preferred, further characterizations

will be performed only on the closed plasma treated chips and the MDA-MB-231 cell line.

Figure 3-10A illustrates live (green) and dead (red) cells within the micro channels of the

chip. This image was taken 14 days after the cells were printed into the microchannels. As seen

in Figure 3-10A, there are an abundant amount of live cells actively growing within the microfluidic

chip. This live/dead investigation confirms that the plasma treated microfluidic chip fabricate from

SU-8 with the DMM, can support cells in a microfluidic environment. The orientation of the cells

(bright green) in Figure 3-10A suggests that the cells are within the microchannels. The spatial

arrangement demonstrated suggests that cells are uniformly distributed within the channels and the

laminar flow of the culture medium within the cells does not affect their attachment or growth.

During printing, external forces act upon cells within the print head (K. Nair et al., 2009).

It is critical that the cells which are printed into the microchannels are not injured or worst, die from

the printing process. To characterize the effects of the printing process onto the cells; a

fluorescence image was taken 24 hours after the cells were printing into the microchannel using a

confocal microscope. Nair et al., stated that cells may be injured during the printing process and

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can recover while others may not. Figure 3-10B confirms that the cytoplasm and nucleus are not

damaged by the printing process and there are no signs of damaged cells within the channels of the

chip. As seen in Figure 3-10B, the nuclei of the cells (brighter green) are well defined. Well

defined nuclei illustrates there are no significant signs of damage onto the nuclei. The lighter green

which represents the cytoplasm appears to start changing its morphology to that of the MDA-MB-

231 cells. Figure 3-10C shows an SEM image of the MDA-MB-231 cells within the microchannel.

This image provides an in-depth view of the cell’s morphology and attachment onto the channel’s

surface. As seen in Figure 3-10C, the cells are well attached onto the surface and its morphology

is that of the MDA-MB-231 cell line.

Figure 3-10. (A) A fluorescence image, taken at 14 days after cells were seeded into the microfluidic chip showing live cell stained green and dead cells stained red. (B) A confocal

image, taken 24 hours after cells were seeded into the microfluidic chips showing the nuclei (stain bright green) and the cytoplasm (stain green) of the cells in the channel. (C) An SEM image,

showing an in-depth view of the cell morphology within the channels.

3.6. Cell Printing and Structural Integrity

Using the multi-nozzle biologics deposition system allows for precise spatial control of

cell placement within the microchannels. Conventional methods of seeding cells to microfluidic

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chips are done by injecting the cells through the inlet of the chip. An investigation was conducted

using the closed chip with the MDA-MB-231 cells to determine which cell seeding method is

better. The results of this investigation are presented in Figure 3-11. According to the data; after

14 days, there are a significant amount of cells present in the chips that used the multi-nozzle

biologics deposition system compared to the chips that had cells injected into chip through the inlet.

Chips with the conventional seeding method had a very slow proliferation rate and a low initial cell

count/attachment. Since cells are injected into the microchannels, there is no control on the spatial

orientation and number of the cells within the channels. Cells may clump together while being

injected; this may cause blockage and uneven distribution of cells. The cell printer resolves these

issues. There is no clumping of cells with the cell printer and cells are uniformly distributed within

the channels. This leads to more initial cells seeded during the seeding process and more cell-cell

interaction for an active proliferation. The exponential cell proliferation trend present in Figure

3-11 confirms the benefits of using the cell printer to deposit cells into the microchannels.

Figure 3-11. The effects of conventional and cell printing seeding methods on cell proliferation within the microfluidic chips.

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Three fluorescence microscopic images were taken are of chips that were seeded with cells

using the cell printer. The three images were taken to give an overview of the entire cell distribution

within the chip. These images were taken 8 hours after cells were printed into the channels. Figure

3-12A is a schematic of the microfluidic chip where the black line illustrates the microchannels.

There are 3 rectangular highlights demonstrating the area in which microscopic images were taken.

Figure 3-12B is a 4x image of the left side of the microchannel, Figure 3-12C is a 4x image of the

center of the microchannel, and Figure 3-12D is a 4x image of the right side of the microchannel.

Cells are pointed out with the arrow. These images confirm the DMM abilities to fabricate cell-

laden microfluidic chips with ease and precision. The uniform distribution of cells within the

microchannels demonstrated the multi-nozzle biologics deposition system’s ability to precisely

place cells within the channels.

Figure 3-12. (A) A schematic of the microchannels on the microfluidic chips. (B) An image of the left side of a microchannels on the microfluidic chip showing the cells (labeled with the arrows) within the channel and channel’s uniformity. (C) An image of the center of a microchannels on

the microfluidic chip showing the cells (labeled with the arrows) within the channel and channel’s uniformity. (D) An image of the right side of a microchannels on the microfluidic chip showing

the cells (labeled with the arrows) within the channel and channel’s uniformity.

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3.7. Conclusions

The DMM system has the capabilities to develop cell-laden microfluidic system. Chips

fabricated with the DMM system have demonstrated their potential to promote cell attachment and

proliferation. Investigations presented in this chapter give way for complex micro-architectural

design for biological applications. The incorporation of a three-dimensional cell printer provided

the added capabilities for precise spatial control of cells within the channels. Spatial orientation of

cell will benefit the fabrication of complex future models. Three-dimensional micro-structures can

be fabricated with the DMM by the layer-by-layer technique. Additionally, the DMM system

provides an economical fabrication technique to produce biological tissue arrays. The data

presented shows that the approach presented in this chapter to fabricate a micro-platform

demonstrates capabilities and potentials to develop cell-laden microfluidic chips.

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CHAPTER 4: INTRODUCTION OF A FREEFORM MICRO-PLASMA SYSTEM FOR THE DEVELOPMENT OF A THREE-DIMENSIONAL CELL-LADEN MICROFLUIDIC CHIP OF IN VITRO DRUG METABOLISM DETECTION

4.1 A Synopsis of Cell-laden Microfluidic Chips

In the field of tissue engineering and regenerative medicine, three-dimensional cell printers

are used to develop tissue scaffolds and building blocks for the generation and regeneration of

functional tissue. The patterning of cells on surfaces is utilized for the development of biosensors,

biomedical devices, and aids in the investigation of fundamental cell biology questions (R. Chang,

Nam, & Sun, 2008; Khalil, Nam, & Sun, 2005; V. Mironov, Boland, Trusk, Forgacs, & Markwald,

2003; W. Sun, Darling, Starly, & Nam, 2004b; Wilson & Boland, 2003). The search for drugs

demands robust and fast methods to find, refine, and test probable drug candidates. Integrating the

advances in the tissue engineering and microfabrication fields creates a potential to develop

biosensors that will produce tissue arrays for pharmaceutical investigations. These sensors can

potentially characterize pathogens, toxicants, odorants, and detect drugs within a given sample(s)

(Aernecke, Snow, Knight, Malliaras, & Tok, 2008; Azad, Akbar, Mhaisalkar, Birkefeld, & Goto,

1992; Bidan, 1992). Most biosensors developed within this integrated field are simplified or

advanced devices that allow for faster and accurate characterization (Sparks et al., 2003). These

sensors do not change the nature of molecular reaction, molecular diffusion, or heat transport

governing laws. The need for a three-dimensional sensor for pharmaceutical investigations is

overwhelming.

Investigations presented in this chapter demonstrate the fabrication of an interconnected

three-dimensional tissue array for pharmaceutical investigations. This platform was developed in

part to function as a biosensor. This sensor can provide a micro three-dimensional environment for

cells to attach, proliferate and differentiate. One unique advantage of this sensor is its laminar fluid

flow within the pores of the chip (Thorsen, Roberts, Arnold, & Quake, 2001). The combination of

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a three-dimensional architecture and laminar fluid flow makes this sensor a prime candidate for

drug characterizations (Thompson et al., 2002). A digital microfabrication device

(photolithographic) is activated to fabricate the internal architecture of sensor. The enclosure of

this biosensor is constructed with the use of micro-molding fabrication technique. All surfaces of

the interconnected architecture are chemically enhanced with the use of a micro freeform dielectric

barrier discharge (DBD) plasma treater. This surface modification provides the appropriate

chemical and mechanical cues necessary for proper cell attachment and proliferation (E.D. Yildirim

et al., 2008; E.D. Yildirim et al., 2010). The surface functionalization illustrated is cell specific

and can be regulated for single or multiple cell (Y. L. Han et al.) studies.

A combination of several tissue engineering and microfabrication techniques was utilized

to develop the sensor presented in this chapter, namely; 1) a digital micro-mirror device, 2) a

freeform micro-plasma system, and 3) a multi-nozzle biological deposition system. 1) The digital

micro-mirror device is a Digital Light Processing (DLP) unit that projects images of ‘.jpeg’,

‘.bitmap’, or ‘.gif’ formats. The micro-mirrors have the option to switch between masks within a

matter of micro-seconds, while offering high resolutions performance in Spatial Light Modulation

(SLM). With ultraviolet (UV) light, this component offers a flexible platform to design and develop

proof of concepts, tissue models, biosensors, and micro-organs. 2) The dielectric barrier discharge

(DBD) technique ignites the plasma which is then delivered through a micro-nozzle. DBDs are

non-equilibrium plasmas operated under atmospheric pressure (Laimer & Störi, 2007). Due to a

non-equilibrium nature, DBD plasmas can generate high energy electrons at cool background gas

temperatures (heavy particles). This unique application of a selective high electron temperature,

and low background temperature enables a rich plasma chemistry in many plasma chemical

processes (Eckstein et al.). Once the micro-plasma is generated, it contacts the surface of

biopolymer and changes the topography and chemistry of the plasma-exposed area. 3) The multi-

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nozzle biologics deposition system is capable of depositing heterogeneous materials, cell types,

and biological factors in a controlled and reproducible manner (R. Chang, Sun, W.,, 2009; W. Sun

et al., 2004a; W. Sun & Lal, 2002a). Cell printing is considered to be an effective tool in the field

of tissue engineering to assemble biologics. This printer executes micron-scale spatial control to

generate cell-laden constructs.

4.2 System Overview

Freeform micro-plasma. The dielectric barrier discharge (DBD) technique generates non-

thermal plasma through a 30 µm micro-nozzle. The micro-plasma is generated by a pulsed power

supply with a variable frequency. Connected to the power supply is the plasma electrode system

with a high voltage electrode coaxially inserted in a dielectric tube of either borosilicate glass or

quartz with a ground electrode wrapped about the outside of the tube. The process gas (or gas

mixture) is purged through the annular gap between the coaxial electrode and the dielectric tube.

When the high voltage electrode is powered, plasma is ignited between the electrodes and a micron-

scale glow-like plasma will appear at the tip of the nozzle. Figure 4-1 presents a flow chart of the

micro-plasma system.

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Figure 4-1. A flow chart of the micro-plasma system.

The plasma generated at the tip of the nozzle is utilized to change the topography and

chemistry of the plasma-exposed area. This phase of the fabrication process is a critical

component in the development of the interconnected tissue array as it is responsible for the

biological enhancement of the chip. Figure 4-2 shows a schematic of the cross-section of the micro-

plasma nozzle treating the surface of a substrate. As seen in Figure 4-2, nano features are created

by the plasma nozzle. The nano features created by the plasma nozzle changes the topology of the

substrate. Combined with the surface chemistry changes, this process would make a hydrophobic

substrate more hydrophilic. In this case, the SU-8 is a hydrophobic substrate and the changes in

the topology and surface chemistry allows for a hydrophilic substrate. This change allows for cells

to attach onto the surface of the channels and actively proliferate. Without the nano features and

the enhanced surface chemistry, cells will not attach onto the channels and the chip would fail.

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Figure 4-2. A schematic showing the cross-section of the micro-plasma nozzle treating the surface of a substrate. The major components of the nozzle are shown along with a photo of a

treated sample is illustrated to demonstrate the effects of the micro-plasma nozzle.

Digital micro-mirroring. The main component of this system is the digital micro-mirroring

device (DMD), an optical semiconductor module that allows for the digital manipulation and

projection of UV light. A computer system is integrated with this microfabrication system to

operate the digital mirrors. The mirrors are mounted directly above the platform and are angled

towards the UV light source. The UV source is adjustable in terms of intensity and exposure time.

The light from the UV lamp travels and uniformly distributes on the micro-mirrors. During the

projection phase, the digital mirrors would either be ‘on’ or ‘off’ depending on the pattern being

projected. Mirrors that are turned ‘on’ would absorb the UV light and project it downwards onto

the substrate, while mirrors that are turned ‘off’ would reflect the UV light in the opposite direction

(T. Nederman et al., 1983). The ‘desired’ light projects pattern downwards towards the platform

and the unwanted light is projected away from the platform. The internal architecture of the chip is

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fabricated using this device. This system is responsible for the fabrication of the internal

architecture of the chip.

Multi-nozzle biologics deposition. This system is inspired by rapid prototyping technology

and is built on a CAD/CAM platform. The biologics printer operates with the cell-friendly

conditions of room temperature and low pressure conditions. This system consists of three motion

arms for three-dimensional spatial control and a material deposition which houses up to four

biological materials at once. The deposition system utilizes micro-valve nozzle systems that can

deposit numerous solutions with a wide range of material and biological properties. The computer

controlled multi-nozzle biologics deposition system eliminates human errors and provides its users

with precision control during fabrication procedures. This printer executes micron-scale spatial

control to generate cell-laden constructs. The final phase of this fabrication process utilizes this

system to precisely deposit cells into the microchannel. With its unique ability of precision and

control, this system enables the user to seed cells into the construct of any complex architecture.

This allows for uniform cell seeding throughout the chip.

4.3 Development of Three-dimensional Interconnected Microfluidic Chips

Fabricated entirely from PDMS, the enclosure of the sensor is developed to house the

internal features. The enclosure comprises of a platform (bottom) and a lid (top). The inlet and

outlet ports are a nylon based luer-lock port (McMaster-Carr, Robbinsville, NJ, USA). PDMS is

mixed at 1:15 ratio, de-gassed and cured in an aluminum mold at 130°C for 10 minutes. The cured

PDMS is cooled and removed from the aluminum mold. This process is repeated for the lid where

the luer-lock ports are placed into position prior to being cured on the hot plate. Figure 4-3

illustrates a model of the PDMS enclosure where a PDMS ring is added for each additional layer.

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Figure 4-3. A Model of the PDMS enclosure of the microfluidic chip.

SU-8 is a chemically amplified, epoxy-based negative photoresist typically used for

producing ultra-thick resist layers during device manufacturing in the semiconductor industry.

However, as a hydrophobic material, the use of SU-8 is limited due to the high degree of non-

specific adsorptions of biomolecules, as well as limited cell attachment (Sant et al., 2011). The

structural integrity and photo-chemical properties of SU-8 makes this material an ideal candidate

for the fabrication of micro-structures. The biological enhancement of SU-8 which allows for cell

attachment, proliferation, and differentiation is presented below. The fabrication of the internal

features starts by pouring and leveling SU-8 within the PDMS slot (bottom of the enclosure). The

bottom enclosure with the SU-8 is soft-baked at 65°C for 20 minutes, then at 90°C for 220 minutes

for stability. Immediately after soft-baking, it is cooled for 30 minutes then exposed at the

recommended exposure time based on the amount of energy required for crosslinking (provided by

the manufacturer). The exposure time with the use of the digital mirrors is 10.75 minutes, a total

exposure of 557 mJ. The exposed sample is then hard baked at 65°C for 15 minutes, then at 90°C

for 30 minutes for structural integrity. Prior to development with the SU-8 Developer (MicroChem

Corp., Newton, MA, USA), samples are cooled for another 30 minutes. During the development

process, all unwanted SU-8 will be washed away. The total development time per sample is 8-15

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minutes. After development, samples are removed and rinse with deionized (DI) water to remove

any excessive materials within the channels.

The second layer follows the same protocol for soft-baking, exposure, hard-baking, and

development. The fabrication of the second layer begins with creating a PDMS enclosure; this

enclosure is as thick as the first layer (500 µm). As listed above, enclosure for the second layer

follows the same fabrication protocol as the fabrication of the bottom enclosure. After the enclosure

is cooled, it is placed on a piece of glass (due to the curing ratio of the PDMS, the PDMS sticks to

the glass without any leaks). SU-8 is then poured and leveled within the enclosure followed by

soft-baking, exposure, hard-baking, and development with the same protocol as the first layer.

Figure 4-4(A) shows a schematic of the fabrication and assembly of the cell-laden microfluidic

chip.

Three chips are featured in this chapter with varying porosity. The width of the channel

directly affects the fluid flow; the wider the channel becomes the more turbulent the flow becomes.

To maintain a laminar flow within the chip, the channels should be as small as applicable to the

chip’s functionality. Hence, the width of should not exceed 1 mm. If the channel’s width exceeds

1 mm its environment can no long be considered a microfluidic environment. With the upper limit

established, a lower limit will present an acceptable range to select three varying pore sizes. A

cell’s diameter ranges from 5 µm up to 50 µm. For cell to have a good cell-cell interaction there

must be multiple cells together. This means the minimum channel width should allow for at least

four cells to attach perpendicular (side by side) to the fluid flow. Hence, the minimum width should

be 200 µm. With the acceptable range being 200 µm to 1000 µm; the three chips selected, features

a low, mid, and upper range microchannels. Several fabricated three-dimensional tissue scaffold

features a pore size of 200 µm to 500 µm (Karageorgiou & Kaplan, 2005; O’Brien, Harley, Yannas,

& Gibson, 2005). The first two chips of pores 300 µm and 500µm will investigate the difference

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of the upper and lower limits of the preferred three-dimensional scaffold pores in a microfluidic

chip. The final chip of 700 µm was selected to investigate the benefits of a larger microchannel

chip. Together, these three chip sample will give a board idea of the cells’ functionality in a three-

dimensional laminar cell-laden microfluidic chip. Figure 4-4(B-D) shows three schematics

illustrating the microchannel orientation of the first and second layers of the microfluidic chip along

with a schematic of the two layers overlapping each other for the 300 µm, 500 µm, and 700 µm

microfluidic chips.

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Figure 4-4. (A) A schematic of the fabrication and assembly of the cell-laden microfluidic chip. (B) A schematic illustrating the microchannel orientation of the first and second layers of the chip

along with a schematic of the two layers overlapping each other for the 300 µm chip, the black bars represents the channel walls and the white bars are the channels. (C) A schematic illustrating the microchannel orientation of the first and second layers of the chip along with a schematic of the two layers overlapping each other for the 300 µm chip, the black bars represents the channel

walls and the white bars are the channels. (D) A schematic illustrating the microchannel orientation of the first and second layers of the chip along with a schematic of the two layers

overlapping each other for the 300 µm chip, the black bars represents the channel walls and the white bars are the channels.

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4.4 Sterilization, Plasma Treatment, and Cell Printing

Sterilization is a critical process whenever samples have to undergo biological

characterization. There are several sterilization methods available; however, the appropriate

method must be chosen in order to prevent any deformation or unwanted changes to the samples.

Since the materials used to fabricate the biosensor are insensitive to high temperatures, the authors

have found that a dry heat sterilization process of 150 °C will suffice.

Prior to cell deposition within the microchannels, all samples are plasma treated with the

in-house freeform micro-plasma. The gas composition used for treatment is 5% oxygen and 95%

helium (C. Wang, Hamid, Snyder, Ayan, & Sun, 2012; E.D. Yildirim et al., 2008; E.D. Yildirim et

al., 2010). The plasma nozzle is programmed to first treat the PDMS (enclosure); plasma treating

the PDMS will create a seal between the PDMS-to-PDMS contact. The second phase of plasma

treatment is treating the SU-8 features for which the micro-nozzle allows precise treatment of the

channels. Immediately after treatment, the Multi-nozzle Biologics Deposition device prints cells

within the channels. Figure 4-5(A) shows a photo of the biologic deposition nozzle printing cells

into the channels of the chip. Once the cells are printed within the first layer of the chip, the second

layer is placed on top of the first layer after being plasma treated (same protocol as the first layer).

The plasma treatment changes the functionalization of the PDMS and SU-8 which creates a seal

between the layers and provides an environment for cells to attach and proliferate. The chip is

complete once the lid is placed on top of the second layer, after cells are deposited within the

channels of the second layer. Figure 4-5(B) shows a photo of a fully fabricated chip, complete with

enclosure, inlet and outlet ports (white), and internal features. Once the cell-laden chip was sealed,

it was immediately placed in the incubator with a fluid line connected to the inlet and outlet of the

chips for a total duration of 14 days where culture medium was perfused continuously through the

chips with the use of a syringe pump at a flow rate of 30 µL/hr. During this time span, several

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biological characterizations are conducted at various time points. Figure 4-5(C) shows a photo of

the incubation period of the chips where the syringe pump perfuse culture medium through the

chips.

Figure 4-5. (A) A photo of the biologic deposition nozzle printing cells into the channels of the chip. (B) A photo of a fully fabricated chip, complete with enclosure, inlet and outlet ports

(white), and internal features. (C) A photo of the incubation period of the chips where the syringe pump perfuse culture medium through the chips.

The working cell suspension that is loaded into the biological nozzle utilized the MDA-

MB-231 cell line. The confluent MDA-MB-231 cells were harvested and counted from 75

cm2 vented flasks using hemocytometer. Cells were then re-suspended to a cell density of 1x106

cells/mL and then loaded into the cell printer where it’s printed into the microchannels. The printing

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process is very short, typically 15-30 seconds per chip. Each print job uses 1 mL of cell suspension.

This prints up to 20 layers with a total print time of at least 5 minutes. With a short print time and

an actively moving cell suspension, the cells in the nozzle does not have sufficient amount of time

to settle. In terms of cell aggregation, cells do gather together. Prior to printing, cells are pipetted

to minimize aggregation. Once the cells are loaded into the microchannels, cell moves around and

aggregate together during their initial attachment phase. The images taken (see the results and

discussions section for images) during the biological investigation does not see the aggregation as

an issue as the cells in the channels appears to be uniformly distributed.

4.5 Cytotoxicity Analysis and Cell Interactions

The first step in ensuring that this interconnected microfluidic chip is a potential

pharmaceutical platform to characterize drugs is to investigate its biological relevance. There are

several biological studies that are presented in this chapter that illustrate strong evidence that this

device is a good sensor for drug investigations. As discussed earlier, there are 3 pore sizes that are

characterized; 300 µm, 500 µm, and 700 µm. Since the architecture is fabricated from photo-

material that is enhanced for biocompatibility, it is important to characterize the cytotoxicity of the

chip. The interactions of the cells within the microchannels are of interest. Healthy cells have the

ability to attach to the substrate and proliferate; this is an indicator that the substrate which the cells

are growing on is not toxic. Biological assays such as fluorometric indicators and Enzyme-Linked

Immunosorbent Assay (ELISA) are utilized to characterize the cytotoxicity of the chips.

Cytotoxicity Analysis was conducted using a live/dead stain where live cells are stained

green and dead cells are stained red. Calcein-AM (Dojindo, Japan, 1 μmol/L) is retained within

the live cells, producing a green fluorescence while cells with damaged membranes allow the

entrance of Propidium Iodide(Sigma-Aldrich, USA, 2 μmol/L) which undergoes a fluorescence

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enhancement upon binding to nucleic acids promoting a red fluorescence in dead cells. Prior to

characterization, chips were washed 3 times with PBS then stained with both reagents and

incubated at 37 ℃ for 15 minutes. After incubation, chips were washed 3 times with PBS to remove

the reagents. The washed chips were then observed under a Laser Scanning Confocal Microscope

（Zeiss 710 META, Germany).

The 300 µm, 500 µm, and 700 µm chips showed similar fluorescence results in terms of

cells growing within the channels of the chips. Figure 4-6(A2, B2, and C2) shows a set of

fluorescence images of live cells stained green using the Live:Dead assay. These images highlight

the live cell’s orientation and uniformity within the channels of each chip. A1 is a fluorescence

image of a channel in the 300 µm pore chip, B2 is a fluorescence image of a channel in the 500 µm

pore chip, and C2 is a fluorescence image of a channel in the 700 µm pore chip. As seen these

images, there are a significant amount of live cells (stained green) growing within the chips, this

trend was observed with all chips and is confirmed by the proliferation study (Figure 4-7). During

this investigation, not much red fluorescence (dead cells) was observed. This is due in part that

when cells die they detaches from the surface on which they were attached to while they were alive

and actively proliferating. Additionally, when these cells die, they tend to float in the culture

medium. During the confocal preparation, samples were washed three times and at this stage

any/all dead cells within the channels were washed away.

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Figure 4-6. (A1, B1, C1) Optical images of a pore of the interconnected chips. The dashed lines highlights the channel walls, the arrow points at cells within the channels. A1 is an optical image of a 300 µm pore chip, B1 is an optical image of a 500 µm pore chip, and C1 is an optical image of a 700 µm pore chip. (A2, B2, C2) are fluorescence images of the live cells stained green with the live dead assay. These images highlight the live cell’s orientation and uniformity within the channels of each chip. A1 is a fluorescence image of a channel in the 300 µm pore chip, B2 is a

fluorescence image of a channel in the 500 µm pore chip, and C2 is a fluorescence image of a channel in the 700 µm pore chip.

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Figure 4-6 (A1, B1, and C1) are optical images of a pore of the interconnected chips. The

dashed lines highlights the channel walls, the arrow points at cells within the channels. A1 is an

optical image of a 300 µm pore chip, B1 is an optical image of a 500 µm pore chip, and C1 is an

optical image of a 700 µm pore chip. Due to the chip’s design and characterization limitations, it

is difficult to capture a full three-dimensional view of the cells within the channels. SEM

characterization can be used to give an in-depth view of the cells within the channels. However,

since the cells are deep inside the enclosure, it would require sectioning the chip to view the inside.

Doing this would destroy the sample. Optical imaging does not interfere with the sample and

presents and idea of what’s happening inside the chip. These optical images present an overview

of the differences within the pores, overlay of the two layers, and the distribution of cell within the

channel. Due to equipment limitations, only one channel can be viewed at once. What’s presented

in the optical images, in terms of cell distribution, is true for both layers throughout the chip’s

architecture.

Cell interactions within the channels play an important role in the development of a fully

functional pharmaceutical sensor (Koh et al., 2008). A fluorometric investigation was conducted

which characterized the cell-cell interaction and proliferation within the fabricated chips. This

characterization was performed with the use of AbD SeroTEC’s Alamar Blue (Ab). The cell-laden

chips were washed with 1x Phosphate buffered saline (PBS) by pumping the PBS through the chips

with a syringe pump at a flow rate of 30 µL/hr. 10% Ab was mixed with culture medium and was

pumped through the chips at 30 µL/hr until the chips were filled with the reagent. The chips were

then disconnected from the syringe pump and were placed in the incubator for 4 hours. After 4

hours, the resulting reagent within the chips were removed from the chips and characterized with a

microplate reader (GENios, TECAN, North Carolina, USA) whose excitation and emission

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wavelengths were 535nm and 590nm respectively. The results from this study are shown in Figure

4-7.

Figure 4-7. Results of the 14 day proliferation investigation of the 300 µm, 500 µm, and 700 µm

microfluidic chips.

All chips in the proliferation study showed an increasing trend of cell proliferation.

According to the data, the chips with the 300 µm pore size had the highest number of cells within

the 14 day study, followed by the 500 µm pore size and the 700 µm pore size chips, respectively.

The high surface area to volume ratio and shear stress within the pores/channels may be credited

for the higher number of cells within the smaller pore sized chip (Karande, Ong, & Agrawal, 2004).

Additionally, at the beginning of the 14 day study, the 700 µm pore chip had the higher number of

cells in its chip. Since the 700 µm chip had the widest channels, a higher volume of cell-suspension

was required to fill the channel during the ‘cell printing’ process. At the end of the study, the 700

µm chip had the lowest cell count and there is a slow progression of growth throughout the

investigation. A lower shear stress and limited cell-to-cell interaction due to wide channels are two

factors that may have contributed to the slowed growth of the 700 µm channel. The 300 µm and

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the 500 µm chips have similar proliferation trends with the only difference being a lower cell count

in the 500 µm chip at the beginning of the study; this may have contributed to the 500 µm chip

having a lower cell number at the end of the study in comparison to the 300 µm chip.

4.6 Drug Metabolism, Cell Morphology and Structural Integrity

To demonstrate effective drug metabolism within the three interconnect chips, a drug is

fed into the chip through the inlet port and metabolized by the cells. Results of this analysis are

used to understand the relative pharmacokinetic efficiency and relevancy interconnected

microfluidics. The drug flow study protocol includes 120 μL of 10 mM EFC (7-ethoxy-4-

trifluoromethyl coumarin) (Sigma-Aldrich) stock solution mixed with 9.88 ml of MDA MB 231

complete cell culture medium. The working solution is perfused into a syringe and infused into the

chips with the syringe pump at a flow rate of 30 µL/hr until the chip is completely filled. Chips are

then incubated with static, non-perfused controls until characterization. During the incubation

period, the drug substrate is metabolized by the cells into the metabolite. This metabolism process

converts the drug 7-Ethoxy-4-(trifluoromethyl)coumarin (EFC) to 7-Hydroxy-4-

(trifluoromethyl)coumarin (HFC) by the enzyme 7-Ethoxycoumarin O-deethylase (R. Chang, Sun,

W.,, 2009; Donato, Jiménez, Castell, & Gómez-Lechón, 2004). Due to characterization limitations,

the cells within the chip cannot be characterized, hence the effluent is characterized to demonstrate

effective drug metabolism. On hours 3, 6, 9, and 12 the effluent is extracted from the chips and is

quantified with a microplate reader (TECAN, GENios). The EFC measured directly correlates to

the HFC within the cell. As the measured EFC decreases, the HFC increases.

After 12 hours there is a trace of the EFC drug concentration found in the 500 µm and 700

µm chips and none in the 300 µm chips. The 300 µm chip demonstrated the sharpest decline of the

EFC drug concentration in the effluent followed by the 500 µm and 700 µm chips, respectively. At

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hour 6, the 700 µm chip had the least amount of EFC drug concentration. This may be due to the

large pores within the chip that allows great diffusion of the drug within the chip. Since equal

amounts of cells were seeded into the chips, the difference of EFC drug concentrations are due to

the pores within the chips. Overall, after 12 hours; cells in the 300 µm chip absorbed the EFC drug

faster compared to the 500 µm and 700 µm. With the exception at the 6 hour mark, cells in the 500

µm and 700 µm chips seem to absorb the EFC drug at the same rate. Figure 4-8 shows the results

of the drug metabolism study.

Figure 4-8. Results of the EFC Drug concentration in the 300 µm, 500 µm, and 700 µm chips over a 12 hours period.

Cell morphology and the internal micro-architecture were evaluated using an FEI/Philips

XL-30 Field Emission Environmental Scanning Electron Microscope (SEM). The images obtained

from the SEM were taken using a beam intensity of 2kV and gaseous secondary electron detectors

of 1.3 Torr. Prior to SEM investigation of the micro-architecture, the appropriate preparation was

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conducted by first sectioning the chip using a sharp straight razor. Since the ion beam cannot

penetrate the PDMS layer, sectioning the chip is the only way the view the internal features. The

method of sectioning does cause some deformation to the internal features by causing the layers to

separate a little. Although the internal features a slightly displaced, the SEM image shown in Figure

4-9A illustrates the internal architecture, interconnectivity, channel formation, porosity, and

structural integrity of the fabricated chips. SEM was conducted on all chips and each chip shown

similar structural integrity (illustrated in Figure 4-9A).

Figure 4-9. (A) A SEM images showing the cross-sectional of a microfluidic chip. This image illustrates the channel’s formation and structural integrity. Each chip showcases the same

formation and structural integrity with their corresponding varying channel width. (B) A SEM image showing the morphology and attachment of the MDA-MB-231 cells within the

microchannel of the chip.

Figure 4-9B is an SEM image taken to illustrate the morphology of the cells within the

chips. The SEM preparation for these samples required the chips to be washed twice with 1x PBS

(pumped through the chips). Then the cells were fixed with 4% glutaraldehyde (Sigma Aldrich,

USA) for 2 hours. After being fixed, each chip was subjected to dehydration by pumping a series

of diluted ethanol (50%, 70%, 90%, 95%, and 100%) through each chip. After the dehydration

process, chips were dried and refrigerated at 4°C for 24 hours. Prior to SEM characterization, each

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chip was sectioned to remove the PDMS layer. As seen in the SEM image, the cells are well

anchored within the chip and showed signs of active proliferation. The morphology differs in

comparison to two-dimensional tissue culture, but is consistent to that of a three-dimensional tissue

culture.

4.7 Fluid Dynamics Computational Analysis

The microfluidic chips presented are not conventional single flow chips. These chips have

interconnected channels and are elevated to 1 mm in total height. The fluids within the chips are

actuated with the use of an external actuator (programmable syringe pump). Microfluidics with an

external actuator is classified as continuous-flow microfluidics. Due to the complex architectural

design and fluid flow of each chip, a computational analysis is needed to illustrate the velocity

gradient and fluid flow.

COMSOL Multiphysics® was used to characterize the fluid flow within the 300 µm, 500

µm, and 700 µm microfluidic chips. COMSOL’s Microfluidic Module is widely used to study

microfluidic devices. The flow of a fluid through a microfluidic channel can be characterized by

Reynolds number using equation 4-1:

𝑅𝑅𝑒𝑒 =𝐿𝐿𝑉𝑉𝑎𝑎𝑎𝑎𝑎𝑎𝜌𝜌𝜇𝜇

4-1

where L is the length scale which equals to 4 times the cross-sectional area over the wetted

perimeter of the channel (4A/P), µ is the viscosity, 𝜌𝜌 is the fluid density, and Vavg is the average

velocity of the flow. Re is often less than 1.0 due to the small dimensions of microchannels. In this

Reynolds number regime, flow is completely laminar and no turbulence occurs.

The boundary conditions for the inlet of each chip are set at the flow rate of the syringe

pump, 30 µL/hr, while the outlet pressure was set at 0 Pa. The cell culture medium was simulated

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through the chips with a density of 1x103 kg/m3 and a dynamic viscosity of 1x10-3 Pas for a period

of one hour. COMSOL’s Microfluidic Module solves the Navier-Stokes equations (equation 4-2):

𝜌𝜌𝜕𝜕𝜕𝜕𝜕𝜕𝜕𝜕

− ∇ ∙ 𝜇𝜇(∇𝜕𝜕 + (∇𝜕𝜕)𝑇𝑇) + 𝜌𝜌𝜕𝜕 ∙ ∇𝜕𝜕 + ∇𝑝𝑝 = 0 4-2

where 𝜌𝜌 denotes density (kg/m3), u is the velocity (m/s), µ denotes dynamic viscosity (Pas), and p

equals pressure (Pa).

Figure 4-10 illustrates the simulation results showing the fluid velocity gradient and the

streamline within the 300 μm, 500 μm, and 700 μm microfluidic chips. The streamline plots are

featured to illustrate the fluid flow throughout the interconnective pores. In each streamline plot,

starting point control was applied with 100 lines originating from the inlet. On the other hand the

velocity gradient plots illustrate fluid direction, magnitude, and most importantly, flow type. Each

microfluidic chip is utilized for their ability to produce laminar flow within its channels. However,

when the architecture is complex, a simulation is beneficial to capture the flow type within the chip.

A set of parallel velocity vector indicates laminar flow, else it is turbulent. The simulation results

are conclusive and states that laminar flow exist in the 300 μm, 500 μm, and 700 μm microfluidic

chips. Additionally, the streamline plot for the 300 μm, 500 μm, and 700 μm microfluidic chips

demonstrate that the fluid is interconnective. These claims are proven correct in Figure 4-10 where

(A1) is a streamline simulation of the fluid flow within the 300 μm interconnected channels. (A2)

is a velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout

the 300 μm microfluidic chip. (A3) is a close-up of the velocity gradient at one of the interconnected

pore within the 300 m microfluidic chip. (B1) is a streamline simulation of the fluid flow within

the 500 μm interconnected channels. (B2) is a velocity gradient showing the magnitude, direction,

and fluid flow type that exist throughout the 500 μm microfluidic chip. (B3) is a close-up of the

velocity gradient at one of the interconnected pore within the 500 µm microfluidic chip. (C1) is a

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streamline simulation of the fluid flow within the 700 μm interconnected channels. (C2) is a

velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout the

700 μm microfluidic chip. (C3) is a close-up of the velocity gradient at one of the interconnected

pore within the 700 µm microfluidic chip.

Figure 4-10. COMSOL Multiphysics simulations illustrating the fluid flow within the 300 μm, 500 μm, and 700 μm microfluidic chips. (A1) is a streamline simulation of the fluid flow within

the 300 μm interconnected channels. (A2) is a velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout the 300 μm microfluidic chip. (A3) is a close-up of the velocity gradient at one of the interconnected pore within the 300 μm microfluidic chip. (B1) is a streamline simulation of the fluid flow within the 500 μm interconnected channels. (B2) is a velocity gradient showing the magnitude, direction, and fluid flow type that exist throughout

the 500 μm microfluidic chip. (B3) is a close-up of the velocity gradient at one of the interconnected pore within the 500 µm microfluidic chip. (C1) is a streamline simulation of the fluid flow within the 700 μm interconnected channels. (C2) is a velocity gradient showing the

magnitude, direction, and fluid flow type that exist throughout the 700 μm microfluidic chip. (C3) is a close-up of the velocity gradient at one of the interconnected pore within the 700 µm

microfluidic chip. Color scale bar unit: µm/s

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4.8 Limitations and Challenges

The digital micro-mirroring device is a fast and an economic approach of fabricating micro-

arrays. One of its greatest limitations is the size of its structures that can be produced. The mirrors

itself is no bigger than 0.7 inches (17.78 mm) in length. With the use of optics the projected pattern

can fabricate larger structures; however, it comes with a loss of resolution. Additional layers are

fabricated by aligning an optical mask onto a physical pattern. As the layer increases it becomes

increasingly difficult to accurately align additional layer. This system has limitations when it

comes to fabricating large micro-structures. Besides size limitations, this device is also limited to

only photo-sensitive materials. The DBD plasma system is idea for localize plasma treatment. As

the treatment area increases, the total treatment time will increase. For large structure it will take

an increasingly long time to complete the treatment process. The biologics deposition system is

similar to the DBD plasma, as they both focuses on developing micro-arrays.

The three-dimensional cell-laden microfluidic chip is a unique product that is developed to

culture cells in a microfluidic environment with directly perfusion with the capability to investigate

drug metabolism (Qudus Hamid, Wang, Zhao, Snyder, & Sun, 2014). Many three-dimensional

tissue scaffolds fabricated culture their cells in a static environment which often lead to a lack of

nutrients getting to the center of the scaffold. Additionally, tissue scaffold tends to be large and

slightly bulky. The architecture of this microfluidic chip is small, interconnected (same as tissue

scaffolds), and most importantly, there is a laminar flow perfusion of culture medium throughout

the chip. The small design allows for only a few thousand of cells to complete seed the array, this

is an economical benefit. This system is advantageous; however, it’s not flawless. Due to its design

these chips can only be used once. Once the chips are sealed they have to be sectioned to gain

access to the inside. The only way to get materials such as cultures and assay to the cells is to pump

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it through it inlet. Additionally, due to its small size, any characterizations conducted will result in

small volumes. This cell-laden microfluidic chip is ideal for investigations where the samples (fluid

characterized) are small and of course, since drugs are expensive; it allows for economic drug

studies. The development of each microfluidic system is specifically designed for a targeted

objective. There are several three-dimensional microfluidic systems that currently exist, each with

its own unique functionality (Bettinger et al., 2005; Chiu et al., 2000; Y. L. Han et al., 2013).

This chapter presents an integrative fabrication technique to develop three-dimensional

interconnected microfluidic tissue arrays for pharmaceutical investigations using digital

microfabrication, biologics printing, and plasma treatment. The architecture of the sensor is

fabricated using digital photolithographic method while the biologics are embedded within the

channels of the internal architecture of chip. Additionally, this chapter illustrates the need for

plasma treatment of the photo-material used to develop the micro-architecture of this chip. This

sensor is proven to be a successful tissue array for pharmaceutical investigation by several

characterization methods. Biological characterizations were performed on each chipset to

demonstrate its potential to host live cells in an interconnected microfluidic chip. The biological

results from the three chipset demonstrated that within the 14 days, cells attached and proliferated

within the interconnected microfluidic environment. Of the three chips studied, the 300 µm chip

had the best cell proliferation, followed by the 500 µm and the 700 µm chip. Additionally, the drug

up-take was slightly better in the 300 µm chips compared to the 500 µm and the 700 µm chips.

SEM characterization were performed to study the cell morphology within the channels, as it is

known that cells in a three-dimensional environment have a different morphology in comparison

to that of two-dimensional cultures; this was proven true with the SEM investigation. Structural

integrity is another focus of this chapter. It is understood that within the micro-features of this

sensor, it would be possible to develop a platform for pharmaceutical investigations. The SEM

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confirms that all chips fabricated are interconnected where the channels are uniformity distributed

within their respective layers. The fabrication procedures and characterizations illustrated in this

chapter demonstrate that the sensor developed in this chapter is a good addition for drug

investigations.

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CHAPTER 5: INTEGRATING THE MULTI-NOZZLE BIOLOGICS DEPOSITION AND MICRO-PLASMA SYSTEMS WITH A FREEFORM

ULTRA-VIOLET HEAD AND A PHOTO-POLYMER MATERIAL DELIVERY SYSTEM TO INVESTIGATE CO-CULTURE OF CANCER CELLS IN A

MICROFLUIDIC ENVIRONMENT

5.1 Feasibility of Testing Protocols, Availability, and Ethical Concerns

The use of animal and human models is limited by the feasibility of testing protocols,

availability, and ethical concerns (Elliott & Yuan, 2011; Parnes et al., 1999) which leads to

monolayer cell cultures being used to investigate potential anti-cancer agents. The issue with

monolayer investigations are that these two-dimensional (2D) models give very little feedback on

the effects of the micro-environment on chemotherapeutic and the heterogeneity of the tumor

(Joseph J Casciari et al., 1994). Cancer progression and invasion into surrounding normal tissue

are influenced through the reciprocal interactions with host stromal cells including fibroblasts,

endothelial cells, and macrophages (Friedl & Alexander, 2011; Nakamura, Matsumoto, Kiritoshi,

Tano, & Nakamura, 1997). Cancer expansion and invasion cannot be studied in this 2D co-culture

model. Additionally, tumor normally expands within a confining environment, which leads to high

stresses in both the tumor tissue and the surrounding tissue. For example, breast adenocarcinoma

cells under compressive strains mimicking the growing cells within a confining environment

showed up-regulation of genes related to invasion and metastasis (Demou, 2010). Thus, it is

important to create a three-dimensional model that recapitulates this confining environment which

hasn’t been realized in current three-dimensional cancer models.

Cancer has long been recognized as many diseases due to its difference among each patient.

In addition to the patient heterogeneity, phenotypic and functional heterogeneity and plasticity

within tumors and between primary tumors and metastases has been brought into tumor

understanding over the past few decades (Bedard, Hansen, Ratain, & Siu, 2013; Burrell,

McGranahan, Bartek, & Swanton, 2013; Junttila & de Sauvage, 2013; Marte, 2013; Meacham &

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Morrison, 2013). One possible cause to the heterogeneity within tumors is the intercellular

genomic instability that leads to a branched evolution in tumors (Burrell et al., 2013). The branched

evolution has now been observed among multiple tumor types, including adenoma-to-carcinoma

transition of the colon (Thirlwell et al., 2010), childhood acute lymphoblastic leukaemia (ALL)

(Anderson et al., 2011), chronic lymphoblastic leukaemia (CLL) (Landau et al., 2013), pancreatic

cancer (Campbell et al., 2010) and breast cancer (Nik-Zainal et al., 2012). Moreover, the

heterogeneity in the micro-environment, that include cancer-associated fibroblast, immune cells,

vascular network and extracellular matrix, is another cause of the heterogeneous hierarchy (Charles

et al., 2010; Junttila & de Sauvage, 2013; Marte, 2013). Those heterogeneities in tumorigenesis

result in the tumor as a complex of distinct subpopulations of tumorigenic cancer cells, their non-

tumorigenic progeny and supporting cells (Figure 5-1). This heterogeneous hierarchy has been

denominated as cancer stem-cell model (Meacham & Morrison, 2013) and has been demonstrated

in various tumor types including acute myeloid leukaemia (AML) (Lapidot et al., 1994), chronic

myeloid leukaemia (CML) (J. C. Y. Wang et al., 1998), breast cancer (Al-Hajj, Wicha, Benito-

Hernandez, Morrison, & Clarke, 2003), glioblastoma (Singh et al., 2004), colorectal cancer

(Dalerba et al., 2007), pancreatic cancer (Li et al., 2007) and ovarian cancer (Curley et al., 2009).

Due to the complexity of such heterogeneity, clinical assessment of anticancer drugs poses to

several practical challenges because of the limitations in current transplantation cancer model

(Bedard et al., 2013). The need for development in preclinical model system essentially increases

as the notion of personalized drugs evolves, which is to choose the efficient drug for each patient

individually (Papillon-Cavanagh et al., 2013).

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Figure 5-1. Adopted from Junttila et al, this schematic illustrates heterogeneity of a cancer model (Junttila & de Sauvage, 2013)

The utilization of the microfabrication techniques to fabricate advanced computing chips

has exponentially increased in the last few decades. Needless to say, this fabrication technique

offers some unique advantages to develop micro-systems. Though many conventional

microfabrication techniques today uses very harsh chemical, the authors believe that the

manipulation of system component and fabrication methods may aided in utilization the

microfabrication techniques used in fabricating computer chip to develop advanced cell-laden

microfluidic systems.

To eliminate the limitations of the traditional mainstays of cancer research (M. Friedrich,

2003), the authors investigates an in vitro three-dimensional cell-laden microfluidic chip which is

developed to co-culture cancer cells. Whether it’s a multicellular spheroid (spherical cell

aggregate), hollow fiber (cell on the outer surface of a hollow cylinder), or multicellular layer

(MCL) (multiple layers of collagen coated semi-permeable support membrane with seeded cells)

models (Bartholomä, Reininger-Mack, Zhang, Thielecke, & Robitzki, 2005; D. Cowan, K. Hicks,

& W. Wilson, 1996; Durand, 1990; R. Durand & P. Olive, 1992; R. E. Durand & P. L. Olive, 1992;

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Elliott & Yuan, 2011; Emfietzoglou et al., 2005; Freyer & Sutherland, 1983; Groebe, Erz, &

Mueller-Klieser, 1994; S. B. Hassan et al., 2001; K. Hicks et al., 1997; K. O. Hicks, Fleming, Siim,

Koch, & Wilson, 1998; K. O. Hicks, Pruijn, Sturman, Denny, & Wilson, 2003; Minchinton, Wendt,

Clow, & Fryer, 1997; Thore Nederman, Helmut Acker, & Jörgen Carlsson, 1983; Nederman,

Carlsson, & Kuoppa, 1988; Nederman, Carlsson, & Malmqvist, 1981; Sutherland & Durand, 1976;

X. Zhang et al., 2005b); the investigated chip serves as a foundation to develop more advance

tissue models.

The fabrication of the cell-laden microfluidic chip presented in this chapter is fabricated

with what is referred to as an integrated solid freeform fabrication system. Because of its targeted

applications; this system eliminates the limitations of conventional photolithography and provides

the end user with the capabilities to develop advantageous three-dimensional models. The

integrated system; 1) eliminates the need for mask by incorporating a dynamic maskless fabrication

technique, 2) allows for direct surface modifications as the model is being fabricated, 3) eliminates

the need for long fabrication processes, 4) eliminates the use of toxic chemicals, 5) allows for

spatially controlled heterogeneous deposition of cells/biologics as the tissue array is being

fabricated. Since the integrated system can develop models on a micro-scale level, this makes

investigations more economic; requiring less reagents, cells, and above all it will allow for

consistency in experimental analysis to due limited interactions with the end user (Hsiao et al.,

2009; P. J. Lee, Gaige, et al., 2007; Ong et al., 2008; Tannock et al., 2002; Toh et al., 2009; Toh et

al., 2005; Tourovskaia et al., 2005; A. P. Wong et al., 2008). The integrated system is specifically

designed for the development of biologically inspired devices, which includes, but is not limited

to, biosensors, lindenmayer systems, and micro-organs.

Presented in this chapter is a fabrication approach in which popular fabrication methods

and techniques are coupled together to develop an integrated system that aids in the fabrication of

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a cell-laden microfluidic system. This system aim to reduce the use of harsh chemical, decrease

the length of fabrication time and enable direct printing of cell as the microfluidic chip is being

fabricated. This chapter illustrates the capabilities, benefits, and challenges of the integrated solid

freeform fabrication system to develop micro-tissue arrays. The biological inspired system can

develop critical three-dimensional cell-laden microfluidic models for the investigation of various

tissue models and biological sensors. A cell-laden co-culture model is presented in this chapter to

demonstrate the system’s capabilities to produce advance functional tissue arrays while studying

cancer cells in a co-culture microfluidic environment. Investigations presented in this chapter

demonstrates; 1) a co-culture of cancer cells in a microfluidic chip, 2) advanced cell printing with

localize surface modification, 3) cell integration, and 4) full additive fabrication of a microfluidic

chip.

5.2 System Integration Analysis

The Integrated Solid Freeform Fabrication System integrates several critical fabrication

components utilized in the fabrication of many biological arrays/platforms. These components

include; three-dimensional spatial control, material deposition, photolithographic, plasma

treatment systems. The three-dimensional spatial control houses all fabrication components on the

z motion arm with connectivity to an x and y arm for a complete three-dimensional motion. The

material deposition component houses the biological nozzle and the photo-polymer nozzle. The

biologics head is a cell-friendly deposition nozzle on the motion arm that is used for the spatial

deposition and orientation of the cells and or biologics into the microchannels. The final nozzle of

the material delivery component is a piston style nozzle which is used to drive material of higher

viscosity, such as photo-polymers. The photolithographic head has a LED ultra-violet (UV) fiber

optic head that is mounted on the motion arm. Photolithographic component is used for the

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crosslinking of the photoresist immediately after deposition. Prior to cell deposition, the plasma

treatment head will treat the channels with a composition of helium and oxygen based plasma.

Figure 5-2 shows an image of the integrated fabrication system with a close-up of the four

fabrication head respectively labeled.

Figure 5-2. (left) an image of the integrated fabrication system, (right) close-up of the four fabrication head respectively labeled.

Three-dimensional spatial control. The three-dimensional spatial control component is

integrated with all components (ultra-violet, plasma, and material delivery nozzles) and functions

independently of each component. Each component of this system has its specific function; if a

function or component is not needed the scripts (program code) can be written to function as

desired. All nozzles/print head are independent of each other and only one head is utilized at once.

All print heads are housed on the third (Z axis) motion arms. All nozzles utilize the spatial

controllers in sequential order to develop and enhance the fabricated arrays while depositing

biologics into the channels. The motion system is controlled by a proportional–integral–derivative

controller (PID controller), this allows for tuning of the entire system to function adequately with

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given any fabrication task. Figure 5-3A shows the orientation of the three-dimensional spatial

control motion arms and its major components.

Freeform ultra-violet micro-nozzle. SU-8 requires direct exposure to UV light for the

development of microchannels. The freeform ultra-violet micro-nozzle has a base LED UV lamp

where a fiber optic cable is fed to the motion head. The fiber optic cable is the placed into the print

head which is specifically designed to filter the light through inter-changeable micro-nozzle. This

component emits a peak UV light of 485 nm through 50 µm to 500 µm nozzles with a

manufacturer’s list maximum exposure of 15 W/cm2. The freeform ultra-violet micro-nozzle is

coupled with the photo-polymer head which allows for immediate crosslinking of the photo-resist

upon deposition. Figure 5-3E shows an image of the freeform ultra-violet micro-nozzle with its

major components.

Photo-polymer head. The photo-polymer head is specifically designed to work with high

viscous material. Due to space limitations, the piston style design of this head enables the author

to drive a small amount of material without requiring a lot of room. This head features a syringe-

pump style deposition system that utilizes standard syringes with inter-changeable nozzles. The

utilization of standard everyday products allows the end-users to work effectively with tissue

culture products. The head is controlled with an Audrino micro-processor which is embedded into

the motion software. The photo-polymer head is coupled with the freeform ultra-violet micro-

nozzle. This placement allows for immediate crosslinking of the photo-resist upon deposition

which retain structural integrity. Figure 5-3B shows an image of the photo-polymer head with its

major components.

Biologics head. The biologics component is inspired by rapid prototyping technology and

is built on CAD/CAM platform, which integrates with the three-dimensional spatial control

component. The biologics printer operates at cell-friendly conditions of room temperature and low

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pressure conditions. Coupled with the spatial control component, the biologics printer can deposit

multiple cell types and bioactive factors in controlled amounts with precise spatial positioning. The

printer utilizes a micro-valve nozzle. This nozzle enables the printer to deposit a wide range of

solutions with a wide range of material and biological properties. This component eliminates

human errors and provides its end users with precision biologics control during fabrication

procedures. The biologics deposition component is capable of depositing heterogeneous materials,

cell types, and biological factors in a controlled and reproducible manner (R. Chang, Sun, W.,,

2009; W. Sun et al., 2004a; W. Sun & Lal, 2002a). Cell printing is considered to be an effective

biofabrication tool to assemble biologics. It will be used as such in this chapter. Figure 5-3D

illustrates a cross-sectional schematic of the biologics head and its major components.

Localized micro-plasma head. The generation of plasma is done by changing the gas types,

flow composition, and applied electric field within the nozzle along with the corresponding process

parameter for generation of the desired ignition. Plasma generation is the excitation of ions that

bombards the substrate’s surface to manipulate its topology, surface chemistry and functional

groups. In this system, the micro-plasma is delivered through the dielectric barrier discharge

(DBD) technique. DBDs are non-equilibrium plasmas operated under atmospheric pressure (Ayan

et al., 2009). Due to a non-equilibrium nature, DBD plasmas can generate high energy electrons at

cool background gas temperatures (heavy particles). This unique character (selective high electron

temperature, and low background temperature) enables rich plasma chemistry in many plasma

chemical processes (Ayan et al., 2008). The micro-plasma component consists of a power supply

and the plasma electrode components. Micro-plasma will be generated by a pulsed power supply

with variable frequency. Connected to the power supply will be the plasma electrode system with

a high voltage electrode coaxially inserted in a dielectric (borosilicate glass or quartz) tube and a

ground electrode wrapped around the tube from the outside. The process gas (or gas mixture) will

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be purged through the annular gap between the coaxial electrode and the dielectric tube. When the

high voltage electrode is powered, plasma ignites between the electrodes and a micron-scale glow-

like plasma will appear at the tip of the nozzle. Once the micro-plasma contacts the surface of

biopolymer, it will change the topography and chemistry of the plasma-exposed area. Depending

on the micro-plasma operation parameters, such as plasma power, gas flow rate, gas composition,

and nozzle tip diameter, the authors can control the surface chemistry and topological features of

the exposed photo-polymer. Figure 5-3C shows a schematic view of the localized micro-plasma

head and its major components. Figure 5-4 presents a flow chart of the integrate system with each

of its five major components outlined with color-coded dashed lines.

Figure 5-3. (A) image of the three-dimensional spatial control system with its major components labeled, (B) an image of the photo-polymer head with its major components labeled, (C) a cross-sectional schematic of the localized micro-plasma head with its major components labeled, (D) a cross-sectional schematic of the biologics head showing its major components, (E) An image of

the freeform ultra-violet micro-nozzle with its major components labeled.

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Figure 5-4. Flow chart of the integrate system with each of its five major components outlined with color-coded dashed lines.

5.3 Manufacturing Methods

The cell-laden microfluidic chip presented in this chapter is fabricated in two parts. The

first part of the chip is referred to the enclosure. The enclosure is fabricated with an aluminum

mold which produces a base enclosure and a lid. The base enclosure has a rectangular slot that is

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later utilized by the fabrication system to print, treat, and deposition cells in the desire micro-

architecture. The lid is a flat and houses the inlet and outlet which is utilized for perfusion

throughout the chips during the incubation period. The enclosure is made primarily out of

Polydimethylsiloxane (PDMS) (Dow Corning, Michigan, USA) and the inlet and outlet ports are

nylon based luer-lock connectors (McMaster-Carr, Robbinsville, NJ, USA). PDMS is mixed at

1:15 ratio, de-gassed and cured in an aluminum mold at 130°C for 10 minutes. Cured PDMS is

cooled and removed from the aluminum mold. This process is repeated for the lid where the luer-

lock ports are placed into position prior to being cured on the hot plate. Figure 5-5B illustrates a

model of the PDMS enclosure. Prior to the fabrication of the internal features of the chip, the

enclosure goes through a dry heat sterilization process of 150 °C for 2 hours.

The second part of the chip is the fabrication of the internal architecture which sits in the

slot of the base enclosure. The internal micro-architecture of the chip is fabricated with the

integrated system using SU-8 2100 (MicroChem Corp., Newton, MA, USA) as the building

material. SU-8 is housed in the photo-polymer print head. The localized micro-plasma head uses

a gas composition of 5% oxygen and 95% helium for plasma treatment (E.D. Yildirim et al., 2008;

E.D. Yildirim et al., 2010). Cells used in studies presented in this chapter are MDA-MB-231 (breast

cancer cells) and HepG2 (Liver cancer cells) (American Type Culture Collection (ATCC)

(Virginia, USA)). Prior to printing, cells are harvested from a 75 cm2 tissue culture flask and

counted then re-suspended at a cell density of 1 x 106 cells/ml (50% MDA-MB-231, 50% HepG2)

in culture medium (50/50, MDA-MB-231 culture medium/HepG2 culture medium) and placed in

the biologics head. The freeform ultra-violet micro-nozzle is set at 100% intensity with a 500 µm

nozzle.

The fabrication of the internal architecture is done in a sequential series of steps. The first

step is the photo-polymer head moving into the slot of the enclosure base, then depositing the SU-

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8 forming a line filament height is 0.5 mm. Once the first filament is printed, the UV head activates

and follow the same toolpath and expose the printed SU-8. This process is repeated until the desire

‘layer’ is printed and exposed (UV). The fabrication method utilized by this three-dimensional

printer is layer-by-layer fabrication. Since the chip has only one layer, the fabrication of a second

layer would be identical to the first (process-wise). Once the microchannels have been created,

the plasma head will activate and move over the path of the printed microchannels, treating them.

Figure 5-5C shows an image of the fabricated microchannels within the slot of the PDMS

enclosure. Once plasma treatment is completed, the biologics head will follow the path of the

microchannels and print cells directly into channels. To seal the chip, the plasma head will activate

once more and treat the PDMS on the enclosure. The plasma treatment allows for a seal on PDMS-

PDMS contact between the lid and the base of the enclosure. After treatment, the lid is placed onto

the base enclosure then incubated. Figure 5-5A is schematic illustrating the fabrication steps of

developing the cell-laden microfluidic chip and Figure 5-5D shows an image of the completed cell-

laden microfluidic chip with the lid and its inlet and outlet ports.

Figure 5-5. (A) a schematic illustrating the fabrication steps of developing the cell-laden

microfluidic chip, (B) a model of the PDMS enclosure, (C) an image of the fabricated microchannels within the slot of the PDMS enclosure, (D) an image of the completed cell-laden

microfluidic chip with the lid and its inlet and outlet ports.

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5.4 Biological Characterizations

Cell labeling. Cells are labeled with Qtracker® Cell labeling kit to track cell in co-culture

within the microfluidics. Qtracker® Cell Labeling Kits are designed for loading cells grown in

culture with highly fluorescent Qdot® nanocrystals. Once inside the cells, Qtracker® labels provide

intense, stable fluorescence that can be traced through several generations, and are not transferred

to adjacent cells in a population. Qtracker® 525 and 625 are used to label the HepG2 and MDA-

MB-231 cell lines, respectively. Qtracker® 525 Emission is 525nm and Excitation is 405-485nm.

Qtracker® 625 Emission is 625nm and Excitation is 405-585nm. Prior to investigation, the

working solutions were made by preparing 10 nM labeling solution, pre-mix 1 μL each of

Qtracker® Component A and Component B in a 1.5 mL micro-centrifuge tube. Incubate for 5

minutes at room temperature then add 0.2 mL of fresh complete growth medium to the tube and

vortex for 30 seconds to complete the working solution. This protocol was followed for both cell

labeling kit for its corresponding cell type. Prior to the cells being loaded into the biologics head,

each cell type was suspended in its corresponding cell labeling working solution for an incubation

period of 45-60 minutes. After incubation, cells were washed twice with complete growth medium

then re-suspended and loaded into the biologics head for printing. A fluorescence microscope and

micro-plate reader (GENios, TECAN, North Carolina, USA) was used for characterization.

Evaluation of cell viability and metabolic activity. Cell viability and metabolic activity is

investigated to assess the changes in function of the cells within the microchannels. The topological

and chemical modification provided by microplasma may induce structural and functional changes

in cellular function. A fluorometric investigation was conducted with the use of AbD SeroTEC’s

Alamar Blue (Ab). The cell-laden chips were washed with 1x Phosphate buffered saline (PBS) by

pumping the PBS through the chips with a syringe pump at a flow rate of 30 µL/hr. 10% Ab was

mixed with the co-culture medium and was pumped through the chips at 30 µL/hr until the chips

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were filled with the reagent. The chips were then disconnected from the syringe pump and were

placed in the incubator for 4 hours. After 4 hours, the resulting reagent within the chips were

removed from the chips and characterized with a microplate reader whose excitation and emission

wavelengths were 535 nm and 590 nm respectively.

Evaluation of cell morphology by microscopic visualization. The morphology of the cells

within the microchannels is visualized by confocal microscopy and Scanning Electron Microscopy

(SEM). Cell morphology and the internal micro-architecture were evaluated using an FEI/Philips

XL-30 Field Emission Environmental Scanning Electron Microscope. The images obtained from

the SEM were taken using a beam intensity of 2kV and gaseous secondary electron detectors of 1.3

Torr. Prior to SEM investigation of the micro-architecture and cell morphology, the lid of the chips

was sectioned using a sharp straight razor. After sectioning, the chip was then prep by first fixing

the cells in 2% Glutaraldehyde (GTA) for 2 hours followed by a dehydration process of submerging

the sample in 50%, 60%, 70%, 80%, 90%, and 100% ethanol in series for 10 minutes. Samples

were stored in a 4 °C refrigerator overnight, then splutter-coated with platinum (approximately 10

nm thick) for visualization.

5.5 System Characterization

Chip fabrication and process parameters. The integrated system is governed by a set of

process parameters. Even though each nozzle functions independently to fabricate the cell-laden

construct, the process parameters of the next process (nozzle) are dependent on the output of the

previous nozzle. The first process in developing a cell-laden microfluidic chip is the deposition of

the structural framework. In this case, it’s the deposition of the SU-8 to build the microchannels.

Deposition of photo-polymers from photo-polymer head is governed by the material viscosity,

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applied pressure, length of the nozzle tip, and the radius of the nozzle. To accurately input the

process parameters needed for a specific filament size, Poiseuille’s Law (Equation 5-1) is utilized.

dV𝑑𝑑𝜕𝜕

=𝜋𝜋8 �

𝑅𝑅4

𝜂𝜂 ��𝑃𝑃𝐿𝐿� 5-1

where R is the radius of the nozzle, P is the applied pressure at the inlet (inner orifice of

the nozzle) of the nozzle, L is the length of the nozzle, and 𝜂𝜂 is the viscosity of the material. The

pressure, P of the photo-polymer head is calculated with Equation 5-2 where F is the applied force

and d is the diameter of the syringe. Experimentally derived; the maximum force at the maximum

feed-rate of 1000 mm/min for a 10 mL syringe of the photo-polymer head is 110 lbs, while the

maximum force at the minimum feed-rate of 1 mm/min of the photo-polymer head is 133 lbs.

𝑃𝑃 =𝐹𝐹

2𝜋𝜋𝑑𝑑2 5-2

The force required for motion is calculated using Equation 5-3, where A is the cross-

sectional area of the syringe, l is the length of total volume of material in the syringe, and v is the

motion speed of the syringe’s plunger. This force is directly proportional to speed, greater forces

will result in faster flow rates.

𝐹𝐹 = 𝜂𝜂𝜂𝜂𝑣𝑣𝑙𝑙 5-3

The motion speed of the syringe’s plunger is calculated using Equation 5-4 where P1 is the

applied pressure and P2 is the pressure at the end of the syringe (where the nozzle is connected), r

is the inner radius of the syringe. The speed at any point is proportional to the change of pressure

per unit length (pressure gradient).

𝑣𝑣 =𝑃𝑃1−𝑃𝑃2

4𝜂𝜂𝐿𝐿𝑟𝑟2 5-4

The second print process is exposing the photo-polymer with the UV head. This process

utilizes the geometry of the fabricated filament in the previous process and then determines the

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optimal motion velocity, 𝑣𝑣𝑢𝑢𝑎𝑎 for the UV head. The motion velocity is calculated using equation

5-5 where, 𝐷𝐷 is the nozzle diameter in mm, 𝜖𝜖 is the exposure energy in mJ/cm2, and 𝜀𝜀 is the system

energy output in mW/cm2. The exposure energy can be looked-up on the photo-polymer’s

manufacturer’s specification sheet. The thickness of the deposition material used to match that on

the manufacturer’s specification sheet. The system energy output is that of the UV light, in this

case it’s 15 W/cm2.

𝑣𝑣𝑢𝑢𝑎𝑎[𝑚𝑚𝑚𝑚/𝑠𝑠] =𝐷𝐷[𝑚𝑚𝑚𝑚]

� 𝜖𝜖[𝑚𝑚𝑚𝑚/𝑐𝑐𝑚𝑚2]𝜀𝜀[𝑚𝑚𝑚𝑚/𝑐𝑐𝑚𝑚2]�

5-5

Unlike the other three print heads, the plasma head does not have the ability to change it

nozzles to any desired size. The plasma head is limited to a 30 µm, 50 µm, 100 µm, and a 500 µm

nozzle. Since the channel widths are 500 µm, the 500 µm nozzle is used for the development of

the cell-laden microfluidic chip. It is a very complex process to predictively model the surface

topology and functional groups with the process parameters required to generate plasma. Yildirim,

et al, has investigated the affect various gas composition and has conclude that the gas composition

of 5% oxygen and 95% helium was best for cell attachment and proliferation (E.D. Yildirim et al.,

2008; E.D. Yildirim et al., 2010). Since plasma treatment is only for the substrate’s surface, the

same velocity (calculated with the used of Equation 5-5) as used with the UV head is used for the

plasma head.

Extensive investigations has been conducted on the biological head that is integrated onto

the system presented in this chapter. Nair used rat adrenal medulla endothelial RAMEC cells

(ATCC, MA) is characterized the biological head. The culture protocol is as recommended by

ATCC. Additionally, Nair quantified live, apoptotic, and necrotic cells as a function of the

mechanical perturbations induced by the process parameters. Nair’s samples printed implementing

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each parameter were treated with the Annexin V staining kit (Biovision, Moutainview, CA.

Manufacturer’s protocol were followed.

Nair’s experimental investigations yielded results in Figures Figure 5-6 through Figure 5-8.

Figure 5-6 indicates the decrease in percentage of live cells with increasing dispensing pressure

and decreasing nozzle tip diameter. Figure 5-7 and Figure 5-8 indicates the increase in percentage

of injured and necrotic cells with increasing dispensing pressure and decreasing nozzle tip diameter.

Nair also reported that the effect of pressure is significantly larger than the effect of the nozzle

diameter while at higher pressures, there is an increase in number of injured cells as well as necrotic

cells. Cell viability varies with dispensing pressure and nozzle diameter. The cell viability

decreases as the pressure increases and the nozzle diameter decreases. The effect of pressure is

significantly larger than the effect of the nozzle diameter.

Figure 5-6. Percentage of live cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008).

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Figure 5-7. Percentage of injured cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008).

Figure 5-8. Percentage of dead cells as a function of dispensing pressure for different nozzle diameters (Kalyani Nair, 2008).

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Nair characterized the process parameters of this printed head and found that it allows for

independent adjustments of the applied pressure, P and nozzle diameter, D and assumes that the

independent variable in Equation 5-6 where, E(y) is the expected value (the mean value) for

percentage of live cells (PL), percentage of injured cells (PI), and percentage of dead cells (PD); x1

and x2 represent the independent variables nozzle diameter and pressure. The constants β0 through

β5 were derived by correlating the experimental data wherein percentage live, apoptotic and dead

cells were determined for a range of process parameters (Kalyani Nair, 2008).

𝐸𝐸(𝑦𝑦) = 𝛽𝛽0 + 𝛽𝛽1𝑥𝑥1 + 𝛽𝛽2𝑥𝑥2 + 𝛽𝛽3𝑥𝑥1𝑥𝑥2 + 𝛽𝛽4𝑥𝑥12 + 𝛽𝛽5𝑥𝑥22 5-6

Nair concluded with the predicted equations of percentage of live (Equation 5-7), apoptotic

(Equation 5-8) and necrotic (Equation 5-9) cells expressed as a function of dispensing pressure and

nozzle diameter.

𝐸𝐸(𝑃𝑃𝐿𝐿) = 0.8563 + 0.655𝑥𝑥1 − 0.0268𝑥𝑥2 + 0.0061𝑥𝑥1𝑥𝑥2 − 0.76𝑥𝑥12+ 0.000352𝑥𝑥22 5-7

𝐸𝐸(𝑃𝑃𝐼𝐼) = 0.037− 0.0469𝑥𝑥1 + 0.00297𝑥𝑥2 − 0.002754𝑥𝑥1𝑥𝑥2 − 0.00003488𝑥𝑥12

+ 0.0283𝑥𝑥22 5-8

𝐸𝐸(𝑃𝑃𝐷𝐷) = 0.099− 0.561𝑥𝑥1 + 0.0242𝑥𝑥2 − 0.00496𝑥𝑥1𝑥𝑥2 + 0.665𝑥𝑥12

− 0.000321𝑥𝑥22 5-9

Nair have developed surface plots for Equations 5-7, 5-8, and 5-9 as a function of the

pressure and nozzle diameters. These plots are presented in Figure 5-9 through Figure 5-11, where

Figure 5-9 is the surface plot for the percentage of live cells as a function of process parameters,

Figure 5-10 is the surface plot for the percentage of dead cells as a function of process parameters,

and Figure 5-11 is the surface plot for the percentage of injured cells as a function of process

parameters.

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Figure 5-9. Surface plot for the percentage of live cells as a function of process parameters (Kalyani Nair, 2008).

Figure 5-10. Surface plot for the percentage of dead cells as a function of process parameters (Kalyani Nair, 2008).

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Figure 5-11. Surface plot for the percentage of injured cells as a function of process parameters (Kalyani Nair, 2008).

Nair’s mathematical model quantifies the effect of the process parameters onto the cells.

If the print head is dynamic or static, equations 5-6 to 5-9 holds true. The final parameter of this

print head to model is its motion velocity. Although the applied pressure is different on this print

head compared to the photo-polymer print head, the mechanism of its motion velocity is the same.

Hence, Equation 5-1 was used to determine the appropriate motion velocity of the biologics print

head.

Utilizing the above process parameters with a 250 µm nozzle on the biologics head and a

500 µm nozzle on the photo-polymer head, UV head, and plasma head; the authors fabricated a

sinusoidal microfluidic chip with a channel width and height of 500 µm each. Since the biologics

head prints inside of the microchannels, the nozzle diameter has to be smaller than the width of the

channel. The 250 µm nozzle allowed for spatial control of the printed cells inside of the 500 µm

microchannels. To check for structural integrity of the fabricated microchannels, an SEM

characterization was conducted after the SU-8 was exposed with the UV head. Figure 5-12 shows

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the results of this investigation. Figure 5-12A shows the uniformity of the channels while Figure

5-12B shows the end of the microchannel in which the direction changes from a horizontal channel

to a vertical channel then back to a horizontal channel. Both SEM images prove that the chip is

that of specified dimensions.

Figure 5-12. (A) SEM image showing the uniformity of the fabricated microchannels, (B) SEM

image showing the end of the microchannel in which the direction changes from a horizontal channel to a vertical channel then back to a horizontal channel.

5.6 Cell integration, Proliferation, and Morphological Investigations

The Qtracker® Cell labeling kit was used track cell in co-culture within the microfluidic

chip. There were two kits that were used to label the cells; the MDA-MB-231 cell line was labeled

with the 625 nm Qtracker® kit while the HepG2 cell line was labeled with the 525 nm Qtracker®

kit. These kits allows for tracking each cell type under a fluorescence microscope and quantitative

characterize each cell type with the use of a microplate reader. Since the both cell lines were mixed

together and printed into the microchannels, it is expected that both cell integrates with each other,

attach and proliferate together. Figure 5-13A is a merged fluorescence image taken through a 525

nm and 625 nm filter showing the MDA-MB-231 cell line in red and the HepG2 cell line in green.

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This image proves that both cell lines are integrated with each other. Figure 5-13B is a phase-

contrast image coupled with Figure 5-13A to illustrate the cell distribution within the

microchannels. Both images were taken at day 7 during the investigation. Cell distribution and

active proliferation throughout the 21 day study was collected, analyzed and is showed in Figure

5-13C. As seen in Figure 5-13C, both cell types have an even cell distribution throughout the chip.

Also, over the 21 days, there is an increase in fluorescence intensity which demonstrates an up-

regulation in cell proliferation for both cells.

Figure 5-13. (A) fluorescence image showing cell distribution and integration of the MDA-MD-

231 cells (red, Qtracker® 625) and the HepG2 cells (green, Qtracker® 525) within the microchannels, (B) a phase-contrast image of the cells in the microchannel, (C) quantitative

results of the cell distribution of the MDA-MB-231 and HepG2 cell lines within the microfluidic chip.

Alamar blue was used as a secondary proliferation characterization method. This will

further confirm the results from the Qtracker® kits. This proliferation study characterizes the total

cell growth within the entire chip. This investigation does not differentiate between cells. For

comparative data, two control chips were investigated where one chip was seeded with only MDA-

MB-231 cells and the other was seeded with HepG2 cells. This study shows the cell proliferation

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within the chips and comparatively show the effects of co-culturing these two cells in a microfluidic

environment. Figure 5-14 shows the results of this study. The data shows that over a 21 day period

under the same microfluidic environment, the MDA-MB-231 cells proliferated the fastest while

the HepG2 cells had the slowest proliferation rate. The chip with both cell lines started with a slow

proliferation rate; however, at the end of the 21 day study, the co-culture chip had higher

fluorescence intensity than the chip with the HepG2 cells. This trend demonstrates that when the

two cells are coupled together, it takes a little longer for them to generate and environment in which

both cell lines can thrived in. After about 7 days, the proliferation trend suggests that the extra-

cellular matrix created by both cell lines allows for an up-regulated cell proliferation trend. The

results from the cell proliferation and the cell tracking/cell integration investigation suggests that

co-culturing of two cell lines in a microfluidic chip with enhanced surface treatment is feasible.

Figure 5-14. Results of the 21 days cell proliferation study of the MDA-MB-231 cell-laden chip

(control 1), HepG2 (control 2) cell-laden chip, and the co-culture (both MDA-MB-231 and HepG2 cell lines) cell-laden chip.

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SEM characterization provides an in-depth look of the cell morphology within the

microchannels. This characterization allows for visualization of cell integration within the

microchannels by their morphologies and confirms that cells are integrated and are growing with

the channels. Figure 5-15 are a set of SEM images showing an overview of the cells within the

microchannels (Figure 5-15A), a close-up of the integrated cells (Figure 5-15B), and the

corresponding morphologies of each cell types (Figure 5-15C and D). As seen in Figure 5-15A

there is a uniform distribution of cells throughout the microchannels. This is the same throughout

the entire chip. This image demonstrates the capabilities of the biological deposition component

of the integrated system to precisely print cells in a controlled pattern. Utilizing the morphologies

of the MDA-MB-231 (Figure 5-15C) and HepG2 (Figure 5-15D) cells lines with the close-up view

of the cells in the microchannel (Figure 5-15B), it is clear that the two cell lines are indeed

integrated throughout the chip.

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Figure 5-15. (A) SEM image showing an overview of the cell distribution within the

microchannel, (B) SEM image showing a close-up of the cells within microchannel, the MDA-MB-231 and HepG2 cells are labeled, (C) SEM image showing the morphology of a MDA-MB-

231 cell, (D) SEM image showing the morphology of a HepG2 cell.

The co-culture of cancerous cells have added benefits. Cancer cells have various stages and can be

found in various organs throughout the body. Studying in details, the integration and migration of

different types and stages of cancer cells can allow for more accurate treatment(s). This

microfluidic device provides a platform for which, at least two types or stages of cancer cells can

proliferate together. Depending on the investigation at hand, researchers can learn about treatment

options, how these cells interact with each other, the differentiation process of one stage to another,

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the morphological changes, and even migration. Granted that this device is in its initial

developmental phase, it has many biological applications in cancer investigations.

5.7 Limitations and Challenges

The fabrication method presented in this chapter is unique and advantageous. While it

allows for fabrication of cell-laden microfluidics without the use of harsh chemical it does have

some limitations. The fabrication process of only utilizing one print head at once and the

microscopic nozzles only allows for the development of micro-systems only. This device cannot

develop systems on the macro scale. It does have the ability to fabricate a series of micro-structures

that can be summed into the macro scale; however, fabricating systems such as these can lead to

prolonged fabrication time. If the systems that are fabricated are without cells, the lengthy

fabrication time is not an issue. However, if the system has cells, the lengthy fabrication time will

decrease the cell viability. As demonstrated in this chapter, the fabrication of one layer does not

affect the cell viability at the end of the fabrication process. However, the introduction of a second

layer will decrease the cell viability due to the use of the UV print head. Fabricating an additional

layer will not lead to the demise of all the cells in the layers below, only a small fraction. The

localize treatment of the UV print head is specifically developed to minimize cell death during the

fabrication process such that the fabrication of advanced micro-structure are possible. In addition

to fabrication limitations, there are characterization limitations. Since the micro-systems are

enclosed in PDMS, depending on the enclosure thickness and microscopy instrument, it may be

difficult to qualitative characterize what’s happening inside of the chip. Characterization

techniques should be considered during the development of advanced micro-systems.

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CHAPTER 6: CONCLUSIONS AND RECOMMENDATIONS

6.1 Summary of the Research

This thesis investigated the integration of maskless fabrication, direct cell deposition, and

surface modification techniques to engineer cell-laden microfluidics. The development of tissue-

on-a-chip, organ-on-a-chip, and body-on-a-chip microfluidics are limited to manufacturing

capabilities and materials. Quiet often material selection is limited to specific additive

manufacturing techniques. Since biological constructs require the use of biologically compatible

materials, there is even more limitations in the development of these constructs. To help resolution

this issue at hand, researchers have demonstrated the advantages of microfluidics and its potential

in life sciences and the development of tissue-on-a-chip, organ-on-a-chip, and body-on-a-chip

platforms. With the listed limitations at hand, the author of this thesis investigated a novel

approach of integrating biologically compatible; additive manufacturing techniques, plasma

chemistry to enhance surface functionalization, direct cell deposition, and manipulation of photo-

polymerization with localized UV exposure to assemble cell-laden microfluidics.

The integrated solid freeform fabrication system eliminates the limitations of conventional

photolithography and provide its end-users with the capabilities to develop advantageous tissue-

on-a-chip, organ-on-a-chip, and body-on-a-chip platforms. The integrated system; 1) eliminates

the need for mask by incorporating a dynamic maskless fabrication technique, 2) allows for direct

surface modifications as the model is being fabricated, 3) eliminates the need for long fabrication

processes, 4) eliminates the use of toxic chemicals, 5) allows for spatially controlled heterogeneous

deposition of cells/biologics as the tissue array is being fabricated. Since the integrated system

can develop models on a micro-scale level, this makes investigations more economic; requiring

less reagents, cells, and above all it will allow for consistency in experimental analysis to due

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limited interactions with the end-user (Hsiao et al., 2009; P. J. Lee, Gaige, et al., 2007; Ong et al.,

2008; Tannock et al., 2002; Toh et al., 2009; Toh et al., 2005; Tourovskaia et al., 2005; A. P. Wong

et al., 2008). The integrated system is specifically designed for the development of biologically

inspired devices, which includes, but is not limited to, biosensors, lindenmayer systems, and micro-

organs. This thesis illustrated the capabilities, benefits, and challenges of the integrated solid

freeform fabrication system to develop cell-laden microfluidics. Several biological investigations

were presented to demonstrate the system’s capabilities to produce advance functional microfluidic

arrays.

Figure 6-1 presents a flow chart illustrating the fabrication processes of the integrated

system to develop a cell-laden microfluidic chip. The fabrication processes outline in Figure 6-1

starts with a PDMS enclosure. The photo-polymer head then moves into position and deposits the

photo-resist to build the micro-architecture of the microfluidic chip. The UV head immediately

exposes the photo-polymer which causes the photo-polymer to change its chemical composition

and retain the desired fabricated micro-architecture. Once the first layer of the micro-array is

completed, the micro-plasma head enhances the surface functional groups within the

microchannels. Cells and biologics are deposited into the microchannel with the use of the

biologics head. If there are multiple layers, this process is repeat. To complete the chip, a PDMS

lid is placed on top of the PDMS base where the chip is then sealed and incubated.

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Figure 6-1. Flow chart illustrating the fabrication process of a cell-laden microfluidic chip using the integrative fabrication process.

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6.2 Research Contributions

The contributions of this research are summarized as follows:

1) The photo-resist, SU-8, is a very popular and frequently used material that is

utilized for microfabrication manufacturing processes. The structural integrity and

well established fabrication protocol used to develop precise micro-structures

permits this material to have boundless potential. The manufacturing processes of

SU-8 to develop micro-arrays is considered to be toxic to cells. Also, bare SU-8

is not biologically compatible. The first research contribution of this research

involves a study of SU-8’s potential to serve as a biologically compatible material

for the development of microfluidic chips with enhanced cell attachment and

proliferation. This study investigated three of the most frequently used surface

modification processes and found that all had some form of biological benefit with

plasma treatment proving to be the better of the three (gelatin, sulfuric acid, and

plasma treatments)

2) The utilization of photo-mask in microfabrication processes is a time consuming

and expensive cost factor. Reducing the utilization of a photo-mask during the

fabrication of micro-arrays allows for the cost effective and faster fabrication

processes. Also, since most photo mask are made from chrome; this adds a

contamination factor for the development of cell-laden microfluidic. The second

contribution of this research is the inspection of utilizing a digital mirroring system

with a multi-nozzle biologics deposition system to assemble cell-laden

microfluidics.

3) The utilization of a three-dimensional cell printer allows for precise cell placement

within the microchannels of the chip. Plasma surface treatment of bare SU-8

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enhances the bio-compatibility of the fabricated microfluidic chip. This research

explored a freeform micro-plasma system for the development of a three-

dimensional cell-laden microfluidic chips. This freeform micro-plasma system

enable its end-users to precisely treat the micro-architecture with the same

accuracy and localization as the cell printer.

4) The development, implementation, and characterization of an additive fabrication

system which utilizes; a multi-nozzle biologics component for precise spatial

printing of cells, a micro-plasma head for localized surface functionalization, an

ultra-violet component for freeform exposure of photo-polymers, and a photo-

polymer material delivery component for direct deposition and fabrication three-

dimensional micro-architecture.

5) The development of a three-dimensional interconnected cell-laden microfluidic

chip to investigate drug metabolism and delivered a chip that produces a

microfluidic environment which facilitates co-culture of cancerous cells. The cell-

laden microfluidic chips have laid a foundation to develop advance tissue-on-a-

chip, organ-on-a-chip, and body-on-a-chip platforms.

6) The integrated system is governed by a set of process parameters. The final

contribution of this research is the development of a numerical model to

characterize and predict each component on the fabrication arm (photo-polymer,

micro-plasma, biologics, and UV heads) of the integrated system.

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6.3 Future Research Recommendations

The work presented in this thesis can be improved upon to include more features and

capabilities in the design and development of cell-laden microfluidics. The following research tasks

have are outlined to be undertaken for future research and development:

1) Dual Functioning Plasma/UV Print Head. The plasma head printed in this thesis

utilize the composition of one or more gases to generate plasma. The gas

composition and the applied voltages determined the output of gases. With that

being said, changing the gas type, composition, and applied voltage, it is possible

to generate UV from the plasma head. The development of a dual function

plasma/UV print head will allow for the elimination of the freeform UV head and

the UV source. This development and implementation allows for a simpler, yet

complex material delivery system with one less micro-controller to program and

integrate. This will allow for faster fabrication time, a smoother integration system

to work with, and a faster transition from UV to plasma, vice-versa.

2) Independent Motion. One of the limitations of the integrated system is its ability

to utilize one material delivery nozzle at a time. This hinders the ability for

instantaneous exposure, surface treatment, and cell deposition. A future

recommendation is the development and implementation where each print head

has its own independent motion arms that are integrated with each other that allows

for instantaneous exposure, surface treatment, and cell deposition. This will allow

for faster fabrication time, the utilization of an extended library of biomaterials,

and the potential of fabricating complex heterogeneous tissue arrays.

3) Cell Mechanics in a Chip – Shear Analysis. While perfusion occurs throughout

the chip, there are mechanical forces that are acting on the cells. Each cell type

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reacts differently to external forces. Biological characterizations have shown that

there are no immediate effects on the cells during the fabrication process, hence

and up-regulated cell proliferation trend. Cell perfusion, external forces, such as

shear forces can alter a cell’s phenotype causing the cell to differentiate. For some

cell types this altering their phenotype is beneficial, for others it’s not. Future

investigation should study the cell mechanics within the cell to understand its

effects on the cells to allow for the development of better cell-laden microfluidics.

4) Tissue/Organs-on-a-Chip and Body-on-a-chip. This thesis has successfully

demonstrates a novel fabrication approach where is it possible to fabricate a cell-

laden microfluidic chip and precise place various cell types into the chip’s

microchannels. The chip studied in this thesis is considered to be a single

microfluidic chip. A body-on-a-chip is the inclusion of several bodily function to

capture the function of a body. This unique fabrication approach allows for the

fabrication of multi microfluidic components on the same platforms. Future

research should investigate the potential of utilizing this fabrication technique to

develop multi-organ and/or body-on-a-chip platforms.

5) Study tumor expansion and invasion. Since this fabrication technique allow for

precise deposition of cells, photo, and surface functionality, a future investigation

can be one in which tumor expansion and invasion is studied. The chip of course

would be a three-dimensional heterogeneous model of a targeted tumor where it

can be imaged using a microscopy devices for characterization. For evaluation of

tumor invasion, the number of cancer cells that infiltrate into surrounding cell layer

can be quantified using an imaging analysis program.

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6) Response to anticancer drug treatment. This thesis has presents a microfluidic

platform that investigated drug metabolism and another platform that studied the

co-culture of cancer cells. Future studies can incorporated the benefits of these

two investigation to understand the response to anticancer drug treatment. This

may lead to the development of a cancerous microfluidic model that enable

researchers to develop pharmaceutical that targets specific cancers.

7) Real-time characterization. Future recommendation would be to develop cell-

laden microfluidics that enable researchers to characterize their samples under

real-time conditions. This model is beneficial in the development of

pharmaceutical products. This platform is developed such that it is connected to a

characterization equipment such as a micro-plate reader, microscope, etc. that

monitors the activities within the chip. This model also enables that study of cell

migration, integration, and invasion which allows for a deeper understanding of

cell under microfluidic environment. Microfluidic chips that allows for real-time

characterization has the potential of developing new consumer products.

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VITA

EDUCATION Ph.D., Mechanical Engineering & Mechanics (Biofabrication) Drexel University 2014 M.S., Mechanical Engineering & Mechanics (Mechanics) Drexel University 2011 B.S., Mechanical Engineering & Mechanics (Thermal Fluids & Sciences) Drexel University 2009

FELLOWSHIPS 2011, 2013 National Science Foundation Summer Institute Short Course Fellowship 2012 National Science Foundation EAPSI for US Graduate Students Fellowship

PATENTS 1. Sun, W., Hamid, Q., “Integratable Assisted Cooling System for Precision Extrusion Deposition in the

Fabrication of 3D Scaffolds”, US 2012/0080814 A1, U.S. Classification: 264/176.1; 165/67. 2. Sun, W., Hamid, Q., Wang, C., “Methods of Generating Ultraviolet Radiation, Plasma- and Ultraviolet-

generating Nozzles, Printing Systems, Method of Generating a Substrate, and Substrates Fabricated According to the Same”, Application # 62003768.

REFERRED JOURNAL ARTICLES 1. Hamid, Q., et al, “Maskless Fabrication of Cell-laden Microfluidic Chips with localized Surface

Functionalization for the Co-culture of Cancer Cells”, Journal of Biotechnology, Status: under review. 2. Hamid, Q., et al, “Fabrication of Biological Microfluidics using a Digital Microfabrication System”, ASME

Journal of Manufacturing Science and Engineering, Status: under review. 3. Hamid, Q., et al, 2014, “Surface Modification of SU-8 for Enhance Cell Attachment and Proliferation within

Microfluidic Chips”, Journal of Biomedical Materials Research Part B: Applied Biomaterials, DOI: 10.1002/jbm.b.33223.

4. Hamid, Q., et al, 2014, “A Three-dimensional Microfluidic Tissue-on-a-chip for Detecting Drug Metabolism”, Biofabrication, 6. DOI:10.1088/1758-5082/6/2/025008.

5. Snyder, J.E., Hunger, P.M., Wang, C., Hamid, Q., Wegst, U.G.K., Sun, W., 2014, “Combined Multi-Nozzle Deposition and Freeze Casting Process to Superimpose Two Porous Networks for Hierarchical 3-Dimensional Microenvironment”, Biofabrication, 6. DOI:10.1088/1758-5082/6/1/015007.

6. Ringeisen, B. R., Pirlo, R. K., Wu, P. K., Boland, T., Huang, Y., Sun, W., Hamid, Q., Chrisey, D.B., 2013, "Cell and organ printing turns 15: Diverse research to commercial transitions" Materials Research Society Bulletin, 38. DOI: http://dx.doi.org/10.1557/mrs.2013.209.

7. Hamid, Q., et al, 2011, “Feasibility of Three-dimensional scaffolds using the precision extrusion deposition with an integrated assisted cooling”, Biofabrication, 3. DOI:10.1088/1758-5082/3/3/034109.

8. Snyder, J.E., Hamid, Q., et al, 2011, “Bioprinting cell-laden matrigel for dual tissue drug metabolism and radioprotection study”, Biofabrication, 3. DOI:10.1088/1758-5082/3/3/034112.

INVITED BOOK CHAPTERS 1. Hamid, Q., et al, 2014, “Computer Aided Tissue Engineering for Modeling and Fabrication of Three-

dimensional Tissue Scaffolds”, Chapter 13, Biomaterials and Regenerative Medicine, edited by Peter Ma. ISBN: 9781107012097.

2. Hamid, Q., et al, 2013, “A Digital Microfabrication Based System for the Fabrication of a Cancerous Tissue Models”, Chapter 9, Biofabrication, edited by Gabor Forgacs and Wei Sun. ISBN: 9781455728527.

SELECTED SCI INDEXED CONFERENCE PROCEEDINGS 1. Hamid, Q., et al, 2012, “Digital microfabrication of tissue arrays for pharmaceutical investigations”, Journal

of Tissue Engineering and Regenerative Medicine, 6, Vienna, Austria. DOI: 10.1002/term.1586. 2. Hamid, Q., et al, 2012, “Fabrication of Micro Organs Using a Digital Micro-Mirroring Microfabrication

System”, ASME/ISCIE 2012 International Symposium on Flexible Automation, St. Louis, MO. DOI:10.1115/ISFA2012-7104.

3. Hamid, Q., et al, 2010, “Precision Extrusion Deposition with Integrated Assisting Cooling to Fabricate 3D Scaffolds”, ASME 2010 Conference on Smart Materials, Adaptive Structures and Intelligent Systems, Philadelphia, PA. DOI:10.1115/SMASIS2010-3804.

4. Hamid, Q., et al, 2010, “Coaxial Electrospinning of Biopolymer with Living Cells”, ASME 2010 First Global Congress on NanoEngineering for Medicine and Biology, Houston, TX. DOI:10.1115/NEMB2010-13282.

http://dx.doi.org/10.1088/1758-5082/6/2/025008

http://dx.doi.org/10.1557/mrs.2013.209

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