Microfluidics for Biological Applications

Microfluidics for Biological Applications Wei-Cheng Tian · Erin Finehout Editors Microfluidics for Biological Applic...

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Microfluidics for Biological Applications

Wei-Cheng Tian · Erin Finehout

Editors

Microfluidics for Biological Applications

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Editors Wei-Cheng Tian General Electric Global Research Center 1 Research Circle Niskayuna, NY 12309

Erin Finehout General Electric Global Research 1 Research Circle Niskayuna, NY 12309

ISBN: 978-0-387-09479-3

e-ISBN: 978-0-387-09480-9

Library of Congress Control Number: 2008930844  2008 Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper springer.com

To my family Erin Finehout

To Gan-Wu, Yu-Hsien, Wei-Hua, Kaitlyn, Darren, and Jennifer Wei-Cheng Tian

Preface

In Nobel Prize winner Richard Feynman’s well-known 1959 speech “There’s Plenty of Room at the Bottom” [1] he marvels that although many biological systems, such as cells, are very small, they are active and perform a number of functions. He then poses the challenge “Consider the possibility that we too can make a thing very small which does what we want – that we can manufacture an object that maneuvers at that level!” [1] In this book we hope to show readers that we are getting closer to meeting this challenge. We have tools to manipulate and analyze small volumes of biomolecules (such as DNA and protein); we can manipulate and analyze individual cells; and we can create nanodrops, the size of a cell, to perform specific chemical reactions. All of these have been made possible by the application of microfluidics. This book consists of a selection of review articles that are intended to show how microfluidics is applied to solve biological problems; why microfluidics continues to play an important role in this field; and what needs to be done next. We will introduce not only the various technologies of microfluidics but also how to link these technologies to different biological applications at the industrial and academic level. Chapters 1-3 give perspective on the history and development of microfluidic technologies. They also serve to give a physical understanding of microfluidic devices. Chapter 1 covers the physics and fluid dynamics of microscale flows. Chapter 2 summarizes the materials and methods used to fabricate microfluidic devices in biological applications. Chapter 3 gives solutions to how these microscale devices can be interfaced with the macro scale world. Chapters 4-6 give overviews on how microfluidic systems have been used to study and manipulate specific classes of components. Microfluidic devices have been used to: prepare, amplify, and analyze DNA samples (Chapter 4); separate and analyze protein mixtures (Chapter 5); and culture, separate, and analyze cells (Chapter 6). Chapters 7-10 focus on specific biological applications of microfluidics: tissue engineering (Chapter 7), high throughput screening (Chapter 8), diagnostics (Chapter 9), and biodefense (Chapter 10). And finally, Chapter 11 discusses

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emerging trends in the microfluidics field and the current challenges to the growth and continuing success of the field. In all the chapters, the authors give information on the biological problems that need to be solved, the current research that is being done to address them, and the obstacles that still remain. In addition, there are summaries of the types of products that have been commercialized in each area. This book is intended to be used at the senior undergraduate or graduate level for students. It will also be a great resource for researchers and scientists in the biotechnology, pharmaceutical, and life science industries. We hope to provide the readers with an overview of microfluidics and its current applications to encourage readers to think about how these technologies could help them in their own fields. Reading through the chapters there are a few recurring themes that merit being mentioned here. The first is that the application of microfluidics isn’t just about saving time, cutting costs, and needing less reagents. Working in the microfluidic regime enables scientists to perform experiments and use techniques that simply aren’t possible at a larger scale. The second theme is that for the microfluidics field as a whole to continue to move forward in the biological area, it is vital that scientists from different fields (engineers, chemists, material scientists, biologist, etc.) work together. Only with such collaborations can one be sure that the right questions are being addressed, the right methods are being applied, and the optimal tools are being used. The editors would like to thank Steven Elliot and Angela DePina at Springer for their help in pulling this book together. We would also like to show our appreciation to the authors for all of the time and effort they put towards writing their chapters. Lastly, we’d like to thank our friends and family for their support and patience during this project.

References: 1. Feynman RP (1960) There’s plenty of room at the bottom: An invitation to enter a new field of physics. Engineering and Science 23:22-36.

Contents

Chapter 1 Introduction to Microfluidics ................................................ 1 Abstract................................................................................................... 1 1 Introduction to Microfluidics............................................................... 2 1.2 History of Microfluidics ................................................................... 3 1.2.1 The beginning: Gas chromatography and capillary electrophoresis.................................................................................... 3 1.2.2 The microfluidic advantage ....................................................... 5 1.2.3 Modular separation, reaction and hybridization systems .......... 7 1.2.4 Integrated systems ..................................................................... 8 1.3 Fluidics and Transport Fundamentals............................................. 10 1.3.1 The continuum approximation................................................. 10 1.3.2 Laminar flow ........................................................................... 10 1.3.3 Diffusion in microfluidic systems ........................................... 12 1.3.4 Surface forces and droplets...................................................... 14 1.3.5 Pumps and valves .................................................................... 16 1.3.6 Electrokinetics ......................................................................... 16 1.3.7 Thermal management .............................................................. 18 1.4 Device Fabrication.......................................................................... 18

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1.4.1 Materials .................................................................................. 19 1.4.2 Fabrication and assembly ........................................................ 20 1.5 Biological Applications .................................................................. 21 1.5.1 Genetic analysis (DNA/RNA) ................................................. 22 1.5.2 Proteomics ............................................................................... 22 1.5.3 Cellular assays ......................................................................... 23 1.5.4 Drug delivery and compatibility.............................................. 24 1.6 The Future....................................................................................... 26 1.6.1 Potential demand/market for microfluidic devices.................. 26 1.6.2 Current products ...................................................................... 27 1.6.3 Challenges and the future ........................................................ 28 References ............................................................................................ 29 Chapter 2 Materials and Microfabrication Processes for Microfluidic Devices ...................................................................................................... 35 Abstract................................................................................................. 35 2.1 Introduction .................................................................................... 36 2.2 Silicon Based Materials .................................................................. 37 2.2.1 Micromachining of silicon....................................................... 39 2.2.2 Bulk micromachining .............................................................. 39 2.2.3 Surface micromachining.......................................................... 46 2.3 Glass Based Materials..................................................................... 49 2.3.1 Microfabrication in glass ......................................................... 51 2.4 Wafer Bonding ............................................................................... 56 2.4.1 Fusion bonding ........................................................................ 57 2.4.2 Anodic bonding ....................................................................... 57 2.4.3 Adhesive bonding.................................................................... 58 2.5 Polymers ......................................................................................... 59 2.5.1 Microfabrication ...................................................................... 59 2.5.2 Polymer materials:................................................................... 64 2.6 Conclusion ...................................................................................... 82 References ............................................................................................ 82 Chapter 3 Interfacing Microfluidic Devices with the Macro World.. 93 Abstract................................................................................................. 93 3.1 Introduction .................................................................................... 94 3.2 Typical Requirements for Microfluidic Interfaces ......................... 94 3.3 Review of Microfluidic Interfaces.................................................. 95 3.3.1 World-to-chip interfaces.......................................................... 95 3.3.2 Chip-to-world interfaces........................................................ 103 3.4. Future Perspectives...................................................................... 112 References .......................................................................................... 113

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Chapter 4 Genetic Analysis in Miniaturized Electrophoresis Systems.....................................................................................................117 Abstract............................................................................................... 117 4.1 Introduction .................................................................................. 118 4.1.1 Status of genetic analyses ...................................................... 118 4.1.2 Genetic analysis by miniaturized electrophoresis system ..... 119 4.2 Microchip Electrophoresis for Genomic Analysis........................ 122 4.2.1 Material and fabrication of electrophoresis microchips ........ 123 4.2.2 Theory of gel electrophoresis of DNA .................................. 125 4.2.3 Gel matrices........................................................................... 126 4.2.4 Novel DNA separation strategies on microchips................... 130 4.2.5 Surface coating methods for microchannel walls.................. 134 4.3 Parallelization in Microchip Electrophoresis................................ 137 4.4 Integration in Microchip Electrophoresis for Genetic Analysis ... 139 4.4.1 Sample preparation on microchip.......................................... 139 4.4.2 System integration ................................................................. 141 4.5 Commercial Microfluidic Instruments for Genetic Analyses....... 144 4.5.1 Commercial microchip electrophoresis instruments for genetic analysis ........................................................................................... 145 4.5.2 Integrated microfluidic instruments for genetic analyses ...... 147 4.6 Microfluidic Markets and Future Perspectives............................. 150 References .......................................................................................... 151 Chapter 5 Microfluidic Systems for Protein Separations ................. 165 Abstract............................................................................................... 165 5.1 Introduction .................................................................................. 166 5.1.1 Advantages of microfluidic chips for protein separations..... 166 5.1.2 Limitations of microfluidic chips in proteomics applications167 5.1.3 Substrates used for proteomic analysis.................................. 167 5.2. Microfluidic Chips for Protein Separation................................... 168 5.2.1 Microchip-based electrophoretic techniques ......................... 169 5.2.2 Microchip chromatography ................................................... 172 5.3 Integrated Analysis in Microchips................................................ 175 5.3.1 Integration of sample preparation with analysis.................... 175 5.3.2 Multi-dimensional separation in microchips ......................... 177 5.3.3 Chips integrated with mass spectrometry .............................. 180 5.4. Future Directions ......................................................................... 180 References .......................................................................................... 181 Chapter 6 Microfluidic Systems for Cellular Applications............... 185 Abstract............................................................................................... 185 6.1 Introduction .................................................................................. 186 6.1.1 Physiological advantages....................................................... 188

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6.1.2 Biological advantages............................................................ 189 6.1.3 Economical advantages ......................................................... 191 6.2 Microfluidic Technology for Cellular Applications ..................... 191 6.2.1 Microfluidic cell isolation/separation .................................... 191 6.2.2 Microfluidic cell culture ........................................................ 200 6.2.3 Microfluidic cell analysis ...................................................... 208 6.3 Commercialization of Microfluidic Technology .......................... 211 6.4 Concluding Remarks .................................................................... 214 References .......................................................................................... 215 Chapter 7 Microfluidic Systems for Engineering Vascularized Tissue Constructs............................................................................................... 223 Abstract:.............................................................................................. 224 7.1 Introduction .................................................................................. 224 7.2 Generating 2D Vascularized Tissue Constructs Using Microfluidic Systems............................................................................................... 226 7.3 Generating 3D Vascularized Tissue Constructs Using Microfluidic Systems............................................................................................... 230 7.4 Hydrogel-based Microfluidic Systems for Generating Vascularized Tissue Constructs................................................................................ 232 7.5 Mathematical Modeling to Optimize the Microfluidic Systems for Generating Vascularized Tissue Constructs ....................................... 235 7.6 Future Challenges ......................................................................... 237 7.7 Conclusions .................................................................................. 237 References .......................................................................................... 237 Chapter 8 High Throughput Screening Using Microfluidics............ 241 Abstract............................................................................................... 241 8.1 Introduction .................................................................................. 242 8.2 Cell-Based Assays ........................................................................ 244 8.2.1 High throughput cell culture.................................................. 245 8.2.2 Cell sorting for high throughput applications........................ 252 8.3 Biochemical Assays...................................................................... 254 8.3.1 PCR ....................................................................................... 254 8.3.2 Electrophoresis ...................................................................... 255 8.3.3 Others .................................................................................... 255 8.4 Drug Screening Applications........................................................ 258 8.5 Users and Developers of µF HTS Platforms ................................ 259 8.5.1 Users: Research labs, academic screening facilities, and pharmaceutical................................................................................ 260 8.5.2 Commercialized products in HTS ......................................... 261 8.6 Conclusion .................................................................................... 262

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8.7 Acknowledgements ...................................................................... 263 References .......................................................................................... 263 Chapter 9 Microfluidic Diagnostic Systems for the Rapid Detection and Quantification of Pathogens .......................................................... 271 Abstract............................................................................................... 271 9.1 Introduction .................................................................................. 272 9.1.1 Infectious pathogens and their prevalence............................. 272 9.1.2 Traditional pathogen detection methods................................ 274 9.1.3 Microfluidic techniques......................................................... 276 9.2 Review of Research ...................................................................... 277 9.2.1. Pathogen detection/quantification techniques based on detecting whole cells ...................................................................... 277 9.2.2 Pathogen detection/quantification techniques based on detecting metabolites released or consumed................................... 294 9.2.3 Pathogen detection/quantification through microfluidic immunoassays and nucleic acid based detection platforms............ 297 9.3 Future Research Directions........................................................... 305 References .......................................................................................... 307 Chapter 10 Microfluidic Applications in Biodefense......................... 323 Abstract............................................................................................... 323 10.1 Introduction ................................................................................ 324 10.2 Biodefense Monitoring ............................................................... 326 10.2.1 Civilian biodefense .............................................................. 326 10.2.2 Military biodefense.............................................................. 328 10.3 Current Biodefense Detection and Identification Methods ........ 330 10.3.1 Laboratory detection............................................................ 331 10.3.2 Field detection ..................................................................... 332 10.4 Microfluidic Challenges for Advanced Biodefense Detection and Identification Methods........................................................................ 333 10.5 Microscale Sample Preparation Methods ................................... 335 10.5.1 Spore disruption................................................................... 336 10.5.2 Pre-separations .................................................................... 336 10.5.3 Nucleic acid purifications.................................................... 337 10.6 Immunomagnetic Separations and Immunoassays ..................... 339 10.6.1 Immunomagnetic separations .............................................. 340 10.6.2 Immunoassays ..................................................................... 341 10.7 Proteomic Approaches................................................................ 345 10.8 Nucleic Acid Amplification and Detection Methods ................. 346 10.8.1 PCR and qPCR detection of pathogens for biodefense ....... 347 10.8.2 Miniaturized and Microfluidic PCR .................................... 348

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10.8.3 Heating and cooling approaches.......................................... 349 10.8.4 Miniaturized PCR and qPCR for biodefense....................... 350 10.8.5 Other Nucleic acid amplification methods .......................... 351 10.9 Microarrays................................................................................. 352 10.9.1 Microarrays and microfluidics............................................. 353 10.10 Microelectrophoresis and Biodefense....................................... 354 10.10.1 Microelectrophoresis technologies .................................... 356 10.11 Integrated lab-on-a-chip systems and biodefense ..................... 358 10.11.1 Full microfluidic integration for biodefense...................... 363 10.12 Summary and Perspectives ....................................................... 363 References .......................................................................................... 365 Chapter 11 Current and Future Trends in Microfluidics within Biotechnology Research ........................................................................ 385 Abstract............................................................................................... 385 11.1 The Past – Exciting Prospects..................................................... 386 11.2 The Present – Kaleidoscope-like Trends .................................... 388 11.2.1 Droplet microfluidics........................................................... 389 11.2.2 Integrating Active Components in Microfluidics ................ 391 11.2.3 Third world - paper microfluidics – George Whitesides ..... 394 11.2.4 Microfluidic solutions for enhancing existing biotechnology platforms......................................................................................... 395 11.2.5 Microfluidics for cell biology – seeing inside the cell with molecular probes ............................................................................ 400 11.2.6 Microfluidics for cell biology – high throughput platforms 401 11.3 The Future – Seamless and Ubiquitous MicroTAS .................... 403 References .......................................................................................... 405 Index........................................................................................................ 413

List of Contributors

Beebe, David J., Ph.D University of Wisconsin–Madison Madison, WI 53706 USA Borenstein, Jeffrey, Ph.D. Draper Laboratory Cambridge, MA 02139 USA Burns, Mark A., Ph.D. University of Michigan Ann Arbor, MI 48109 USA Chang, Dustin S. University of Michigan Ann Arbor, MI 48109 USA Chang, Hsueh-Chia, Ph.D. Center for Microfluidics & Medical Diagnostics Notre Dame, IN 46556 USA University of Notre Dame Notre Dame, IN 46556 USA Chung, Yao-Kuang University of Michigan Ann Arbor, MI 48109 USA

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Cropek, Donald, Ph.D. U.S. Army Corps of Engineers Champaign, IL 61822 USA Du, Yanan, Ph.D. Massachusetts Institute of Technology Cambridge, MA 02139 USA Harvard Medical School Cambridge, MA 02139 USA Gordon, Jason E., Ph.D. Midwest Research Institute Kansas City, MO 64110 USA Horn, Joanne, Ph.D. Microchip Biotechnologies Inc. 6693 Sierra Lane, Suite F, Dublin, CA 94568 USA Jain, Akshat University of Michigan Ann Arbor, MI 48109 USA Jovanovich, Stevan, Ph.D. Microchip Biotechnologies Inc. 6693 Sierra Lane, Suite F, Dublin, CA 94568 USA Khademhosseini, Ali, Ph.D. Massachusetts Institute of Technology Cambridge, MA 02139 USA Harvard Medical School Cambridge, MA 02139 USA

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Kuo, Chuan-Hsien University of Michigan Ann Arbor, MI 48109 USA Langelier, Sean M. University of Michigan Ann Arbor, MI 48109 USA Lee, Abraham P., Ph.D. University of California at Irvine Irvine, CA 92697 USA Micro/nano Fluidics Fundamental Focus (MF3) Center Irvine, CA 92697 USA Lin, Gisela, Ph.D. Micro/nano Fluidics Fundamental Focus (MF3) Center Irvine, CA 92697 USA Mofrad, Mohammad R. Kaazempur, Ph.D. University of California at Berkeley Berkeley, CA 94720 USA Noori, Arash McMaster University Hamilton ON L8S 4L7 CANADA Park, Jihyang University of Michigan Ann Arbor, MI 48109 USA Puccinelli, John P. University of Wisconsin–Madison Madison, WI 53706 USA

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Rhee, Minsoung University of Michigan Ann Arbor, MI 48109 USA Selvanganapathy, P. Ravi, Ph.D. McMaster University Hamilton ON L8S 4L7 CANADA Sengupta, Shramik, Ph.D. University of Missouri Columbia, MO 65211 USA Shaikh, Kashan A., Ph.D. GE Global Research 1 Research Circle, Niskayuna NY 12309 USA Sommer, Greg J., Ph.D University of Michigan Ann Arbor, MI 48109 USA Takayama, Shuichi, Ph.D. University of Michigan Ann Arbor, MI 48109 USA Tavana, Hossein, Ph.D. University of Michigan Ann Arbor, MI 48109 USA Upadhyaya, Sarvesh McMaster University Hamilton ON L8S 4L7 CANADA

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Wang, Fang University of Michigan Ann Arbor, MI 48109 USA Wang, Hong, Ph.D. Louisiana State University Baton Rouge, LA 70803 USA Wang, Xuefeng, Ph.D. GE Global Research 1 Research Circle, Niskayuna NY 12309 USA Weinberg, Eli J., Ph.D. Draper Laboratory Cambridge, MA 02139 USA Zeitoun, Ramsey I. University of Michigan Ann Arbor, MI 48109 USA Zhu, Li, Ph.D. GE Global Research 1 Research Circle, Niskayuna NY 12309 USA

Chapter 1 Introduction to Microfluidics

Greg J. Sommer, Dustin S. Chang, Akshat Jain, Sean M. Langelier, Jihyang Park, Minsoung Rhee, Fang Wang, Ramsey I. Zeitoun, and Mark A. Burns University of Michigan, Ann Arbor, MI 48109 Correspondence should be addressed to: Mark Burns ([email protected])

Keywords: microfluidics, history, fundamentals, applications, commercialization

Abstract Microfluidics – the manipulation and analysis of minute volumes of fluid has emerged as a powerful technology with many established and relevant applications within the biological sciences. Over three decades of research has yielded a wealth of techniques for improving biological assays through both the miniaturization of existing methods, as well as the development of novel analytical approaches. In this introductory chapter we provide an overview of microfluidic technology, beginning with a historical look at the field’s origins. We also present brief synopses of the fundamental physical phenomena driving microfluidics, the techniques employed in device fabrication, and biological applications that have benefited from microfluidic implementation. Finally we conclude with an outlook to the fu-

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ture of the field as microfluidic technology shifts from research laboratories into commercial ventures. This chapter is meant to familiarize readers who may be new to the field of microfluidics, while highlighting areas that will be explored in more detail throughout the text.

1 Introduction to Microfluidics Microfluidic technology has evolved over the past few decades from a molecular analysis endeavor aimed at enhancing separation performance through reduced dimensions, into a diverse field influencing an everexpanding range of disciplines. Microfluidic techniques are being employed in chemistry, biology, genomics, proteomics, pharmaceuticals, biodefense, and other areas where its inherent advantages trump standard methodologies. From a biological standpoint, microfluidics seems especially relevant considering that most biological processes involve small-scale fluidic transport at some point. Examples stem from molecular transfer across cellular membranes, to oxygen diffusivity through the lungs, to blood flow through microscale arterial networks. Microfluidics can also provide more realistic in vitro environments for small-scale biological species of interest. Figure 1.1 provides comparative length scales for several biological structures, as well as common micro-fabrication structures used in microfluidic and MEMS technology.

Fig. 1.1 Approximate length scales for several biological and micro-fabrication structures.

Introduction to Microfluidics

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In this chapter we will provide a brief introduction to microfluidics, beginning with a historical perspective of the field’s origins and concluding with an outlook on the future. We also present some fundamental transport principles, materials and fabrication basics, and specific biological applications with the aim of familiarizing the reader with important microfluidicrelated concepts and highlighting areas that will be explored further throughout the text.

1.2 History of Microfluidics 1.2.1 The beginning: Gas chromatography and capillary electrophoresis The mid-20th century saw explosive growth in the applicability of chromatography: a technique that revolutionized the field of separation sciences by exploiting molecular distributions between mobile and stationary phases within a column. Theoretical work by Golay [1] on gas chromatography (GC) and van Deemter [2] on liquid chromatography established scaling arguments showing that improved performance could be achieved by reducing open column diameters and packed column particle sizes. Thus columns began being fabricated from fused silica capillaries with diameters on the order of micrometers. Around the same time, capillary electrophoresis (CE) was gaining popularity as a method to separate charged biomolecules. Here too, small bore capillaries proved advantageous as the larger surface area-to-volume ratio allowed for higher applied electric fields and, therefore, improved separation performance. But the roots of microfluidics truly lie in the microelectronics industry. As chemists and biologists were searching for means to further miniaturize their analytical methods, the microelectronics industry was improving its silicon-based micromachining processes using photolithography, etching, and bonding techniques [3]. The merging of the bioanalytical and microelectronics disciplines can be considered the birth of microfluidics. The first silicon-based analysis system was published in 1979, in which Terry et al. from Stanford University fabricated a miniature GC air analyzer on a silicon wafer [4] (Fig. 1.2). However, it was the seminal works by Manz and others in the early 1990’s that demonstrated the microfluidic potential for addressing issues facing analytical methods, and spawned the term micro Total Analysis Systems (µTAS) [5-7].

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Fig. 1.2 Miniature gas chromatograph developed by Terry et al on a silicon wafer in 1979. [8] - Reproduced by permission of The Royal Society of Chemistry.

While researchers continued miniaturizing gas and liquid chromatography columns [6, 9], many of the first successful microfluidic separation devices employed electrophoretic techniques due to the relative simplicity of applying an electric potential to a microchannel versus a high-pressure source such as those required for high pressure liquid chromatography (HPLC). In 1992, Manz et al. demonstrated the first on-chip CE system and initiated the new microfluidic era in separation sciences. That same year, Mathies et al.proposed high-throughput electrophoretic sequencing on arrays of microfluidic devices [10]. In 1993, Harrison et al. demonstrated a micro-CE system in glass which could separate amino acids with about 75,000 theoretical plates in 15 seconds [11]. The next year, Woolley and Mathies successfully miniaturized a microfluidic capillary gel electrophoresis system for DNA analysis, which boasted separation times in as little as 120 seconds [12]. The microfluidics boom had begun. The mid 1990s brought many new concepts and devices as researchers began to investigate microfluidic uses for not only separations, but other applications as well. So why all the ruckus? In the next section we will highlight some advantages that microfluidics can provide over conventional, macroscale methodologies.


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1.2.2 The microfluidic advantage Silicon micromachining enabled fabrication of channels and features with precision on the order of 1µm. This technological feat enabled the manipulation of micro- (10-6) to atto- (10-18) liter volumes of fluid. Such control brings several advantages from both analytical and economic viewpoints. Here we briefly outline those advantages, while noting that many of these concepts will be further explored throughout this chapter and text. A summary of the advantages are listed in Table 1.1. Table 1.1 Summary of advantages attained with microfluidic systems Microfluidic Advantage Description Less sample and reagent Microfluidic devices typically require 102 – 103 less sample volume than conventional assays. consumption Enhanced heat transfer

Higher surface area-to-volume ratio of microfluidic channels increases effective thermal dissipation.

Faster separations

Higher E-fields results in faster sample migration.

Laminar flow

Low Reynolds number flows reduce sample dispersion.

Electrokinetic manipulation

Electroosmotic flow enables fluid pumping with flat "plug-like" velocity profiles solely via applied E-fields.

Lower power consumption

Fewer components and enhanced thermal dissipation require less power input.

Parallelization

Several assays can be “multiplexed”, or run in parallel on a single chip.

Portability

System integration and reduced power allows for assays to be conducted using portable, hand-held device.

Improved separation efficiency

Efficiency in electrophoretic and chromatographic separations (i.e. number of theoretical plates) proportional to L/d.

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Several measures of analytical performance can be improved through miniaturization. Perhaps one of the most obvious advantages of smaller channel sizes is reduced reagent consumption, leading to less waste and more efficient assays. Reduced reagent consumption becomes especially advantageous for many biological applications where reagents can be very expensive (e.g. antibodies), and sample volumes are often limited. Additionally, the separation efficiency (i.e. number of theoretical plates) of chromatographic and electrophoretic systems is proportional to L/d: the length of the separation channel over its diameter. Therefore long and narrow channels enable improved peak-peak resolution. Because they are so narrow, microfluidic channels also boast flows with very low Reynolds numbers: often Re < 1, meaning the flow is laminar. Such laminar flows inhibit additional dispersion from affecting the band width of a separated plug. Diffusion, however, is more prominent at smaller scales and can be advantageous for mixing applications where, despite very laminar flow, mixing can occur solely via diffusion. Narrow channels also dissipate heat more efficiently, allowing for higher electric fields in electrophoretic systems without adverse Joule heating effects on separation efficiency. As a result the assays will require less time as higher electric fields lead to faster separations. Microfluidic devices often achieve fluid transport with few mechanical components, which can significantly reduce an assay’s complexity and power consumption over its macroscale counterpart. For example, electroosmosis is a process in which bulk electrolytic fluid in a channel is dragged via viscosity by migrating ions near an inherently charged channel wall under the application of an electric field. Electroosmosis allows for: a) bulk fluid “pumping” using only electric fields, thereby eliminating any moving parts, and b) a plug-like, non-parabolic fluid velocity profile that eliminates dispersion caused by parabolic pressure-driven flow. Bulk fluid transport has also been demonstrated on microfluidic devices using other pumping techniques such as capillary wicking, evaporation, thermal gradients, and chemically-induced flow. With many conventional assays it is possible to integrate all analytical steps (sample loading, rinsing, reactions, separation, detection, etc.) into a single, fully-automated platform. Such integration reduces necessary human involvement, potential environmental contamination, and analysis time. With µTAS, or lab-on-a-chip (LOC) devices, such integration can greatly reduce the cost per analysis while providing high throughput through parallelized or multiplexed devices. They can also be potentially integrated into a portable, hand-held format for a variety of point-of-care


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(POC) applications where proper laboratory access is not available or rapid analysis time is required, including bedside patient care, military and border patrol, and global healthcare scenarios. As fabrication procedures become more standardized, the cost per chip will decrease enabling the production of inexpensive, single-use, disposable chips.

1.2.3 Modular separation, reaction and hybridization systems Along with the initial demonstrations of microfluidic separation systems in the early 1990’s, researchers began exploring other methods with which to fill the “analytical toolbox” necessary to build the envisioned integrated systems. Further motivation for the microfluidics community arose from the explosion of genomics in the 1990’s. Biologists were increasingly interested in exploring DNA and decoding genes and chromosomes, as evidenced by the appeal and success of the Human Genome Project. Therefore much of the early research was directed towards DNA amplification, hybridization and sequencing. One technology that naturally found its way into microfluidic devices was that of microarrays. Microarrays, in which minute biological samples are immobilized as individual spots that may hybridize with an introduced sample, allow for extremely large numbers of parameters to be screened at any one time. In 1991, the use of standard lithography to pattern an array was first introduced by Fodor et al. [13] (Fig. 1.3). Since then, microfabricated arrays have found a home using microfluidic technologies. Microarrays have been developed to pack a maximum amount of DNA strands into a minimal amount of space. As early as 1995, a 96 microwell array was used to detect an organism’s gene expressions [14]. A year later, a DNAchip which had 1046 different DNA strands was developed demonstrating the potential power of microfabricating DNA analysis devices [15].

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Fig. 1.3 DNA microarray fabricated lithographically by Fodor et al in 1991. Each square represents one gene sequence and is 50µm wide (Reprinted from [13] with permission from AAAS.)

Early microfluidic reactors were also primarily focused on biological applications. Aside from relatively simple reactions within microarrays, an important biological reaction that benefited from miniaturized formats is the polymerase chain reaction (PCR). PCR is a technique that uses enzymatic transcription to systematically amplify parts or all of a DNA strand, triggered via thermal fluctuations. On-chip PCR incorporation was made possible due to rapid and efficient heating and cooling of extremely small sample volumes, allowing for quick and proficient thermocycling. In 1995, Northrup et al. developed the first microfabricated device capable of thermocycling and PCR reactions [16]. On-chip PCR would prove to be instrumental in future DNA sequencing and genotyping devices.

1.2.4 Integrated systems Modular system integration is particularly advantageous in µTAS devices for several reasons. First, the sample can be isolated from the outside environment, reducing error caused from human contact and sample contamination. In addition, having all processes located on a single chip can, in theory, reduce sample-processing time and allow for a fully-automated analysis. This potential has prompted the idea of integrated devices for point-of-care diagnostics and clinical analysis. However, full integration of numerous modular components is not a trivial task and requires the resolution of numerous problems, including sample injection, pumping, data dissemination and product retrieval.


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At the onset of modular device development, integrated systems were generally simple and relatively crude. Technological efforts were focused on designing simplistic systems that performed one or two actions instead of many. One of the first integrated DNA analysis systems was presented by Burns et al. in 1998 (Fig. 1.4). This device was engineered to meter a precise DNA sample volume, mix it with reagents, amplify it, separate it using gel electrophoresis, and detect the fluorescence signal on-chip [17]. Since that time, DNA separation stages have also been equipped with both on-chip fluorescent and conductivity detectors for on-chip analysis. However, a problem plaguing many of these systems, from a diagnostic standpoint, is that they lack an easy and reliable way to process raw sample, such as blood or saliva.

Fig. 1.4 Integrated DNA analysis device developed by Burns et al. in 1998 (Reprinted from [17] with permission from AAAS.).

During its infancy, microfluidics exhibited the potential to explode into a field rich with powerful applications and commercial successes. Some likened its promising impact to that of the integrated circuit (IC) industry. Today, however, the microfluidics field is arguably not where it was envisioned ten years ago. So what factors have impeded to the field’s emergence as a prevailing technology? We will further explore that question later on in the chapter, but for now we remind the reader that microfluidics continues to be a vibrant and active research area with researchers tackling the challenges hindering its widespread acceptance.

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1.3 Fluidics and Transport Fundamentals Researchers employ many fundamental transport and scaling principles for manipulating and analyzing microfluidic flows. In this section we introduce some of the important concepts governing microscale transport, with the hope of elucidating several advantages of microfluidics over macroscale techniques. We also introduce several dimensionless groups (Table 1.2) that can be used to evaluate the relative importance of different phenomena.

1.3.1 The continuum approximation Unlike solids, fluids (especially gases) consist of molecules that are reasonably widely separated. However, in fluid mechanics, despite the fact that properties like velocity and density vary wildly at the molecular scale, we usually view fluids as “continuous” and discuss “average” fluid properties rather than considering the properties of each molecule. Does this continuum approximation still hold in microfluidics, where fluid volumes are very small? Indeed, for the volumes typically encountered in microfluidics, the continuum approximation remains sufficiently valid. A 1 picoliter [(10 µm)3] volume of fluid still contains 3 x 1013 water molecules, large enough for us to consider their average rather than individual behavior. Typically, for most properties the continuum approximation does not break down until we approach length scales on the order of several molecular diameters [18]. The continuum approximation is important because it allows us to analyze microfluidic flows with the same governing principles developed for macroscale fluid mechanics. However, as researchers continue to drive fluids into smaller length scales (i.e. nanofluidics and beyond), the continuum approximation starts to break down and we must develop new approaches for analyzing these small regimes.

1.3.2 Laminar flow Microfluidic devices almost always boast smooth laminar flow, as opposed to turbulent flow which is of a stochastic nature and marked by the presence of “eddies” that disrupt parallel streamlines. The Reynolds number


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(Re) is a dimensionless parameter used to determine the transition from laminar to turbulent regimes, with Re < 2100 considered laminar for flow in cylindrical channels. The Reynolds number for this flow is defined as

Re =

ud

(1.1)

ν

where u is the flow velocity, ν is the kinematic viscosity, and d is the channel diameter. In fluid mechanics terms, the Reynolds number compares the magnitudes of inertial force to viscous forces in a flow. Because Re ∝ d , the small dimensions of microfluidic channels are responsible for very low Reynolds numbers, resulting in laminar flows. In fact, for most microfluidic applications Re < 1. Since microfluidic flows are usually laminar, simple flows like Poiseuille flow are commonly encountered. Poiseuille flow occurs when we have steady, fully-developed pressure driven flow of a Newtonian fluid in a channel. The velocity profile for Poiseuille flow is parabolic, with the maximum velocity being in the center of the channel. The equations for Poiseuille flow in a cylindrical channel are as follows [19]

r v z (r ) = 2U [1 − ( ) 2 ] R R 2 dP 8µ dz

(1.3)

πR 4 dP 8µ dz

(1.4)

U =−

Q=−

(1.2)

where v z is the velocity of the fluid along the axis, U is the mean velocity (maximum velocity at centerline of channel = 2U), R is the radius of the channel, r is the radial distance from the axis, z is the distance along the axis, P is the pressure along the channel, µ is the fluid viscosity and Q is the volumetric flow rate. While laminar flow is advantageous in many microfluidic applications (i.e. in electrophoretic separations for reducing band broadening due to dispersion, or in diffusion-based separation devices such as the H-filter), it can also be a nuisance, such as in processes where mixing is necessary. Most chemical and biological assays require mixing, or dilution, at some point.

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Several clever techniques have been developed to achieve mixing in laminar regimes, mostly through geometric design or by taking advantage of enhanced diffusivity across small dimensions.

1.3.3 Diffusion in microfluidic systems Diffusion – the stochastic process by which molecules drift from one region to another – is another property that takes on increasing importance in microfluidic systems as channel dimensions are reduced. Diffusive transport is driven by random thermal motion of particles such that, given enough time and the absence of external influences, a species will be homogeneously distributed throughout a finite volume. Most microfluidic systems combine diffusive transport with convective flow of the bulk fluid; therefore it is helpful to compare the relative importance of each effect. The Sherwood number, a dimensionless number representing the ratio of convective mass transfer to diffusive mass transfer in a system, is defined as

Sh =

kd D

(1.5)

where k is the mass transfer coefficient, d is the characteristic length of the system (e.g. channel diameter), and D is the diffusion coefficient. For most macroscale systems, Sh is large, resulting in convective transport being dominant over diffusive transport. However, for microfluidic systems, the Sherwood number is much lower due to the presence of the characteristic length scale in the numerator. Therefore, diffusion assumes much more importance in microfluidic systems. Diffusive transport without flow (or perpendicular to streamlines with flow) can be estimated from the simplified mass transfer equation:

∂ 2C ∂C =D 2 ∂x ∂t

(1.6)

Consider a stationary liquid system with a step change in species concentration such that the concentration is held at C* for all time t at a certain location. Let there be no other source for the substance in the liquid and no other mode of transport besides diffusion. At time t, the concentration will be C*/2 at a distance of about

Dt from the source, and will be 1% of C*


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at a distance of about 4 Dt (Fig. 1.5). Therefore, diffusion has a significant effect up to a distance of about 4 Dt . For polymers or proteins in a fluid, D is on the order of ~10-7 cm2/s. So for a time period of about 10 seconds, the distance at which diffusion is significant will be about 40µm, a length that is usually negligible for macroscale purposes but can be a significant distance in microfluidic systems. Note that, at twice that distance (i.e. 8 Dt ), the concentration is only 10-8C*.

Fig. 1.5 Diffusion-dependent concentration profile at time t for a solute with diffusion coefficient D.

Several mixing and separation techniques have been developed exploiting this diffusive effect in microfluidic systems. Readers are especially referred to the H-filter [20] developed by Paul Yager’s lab at the University of Washington for an example of an efficient and highly-effective device that uses this very simple concept (Fig. 1.6). Here, species in a stream are separated into two diverging flows based on their differing diffusivities across a microchannel.

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Fig. 1.6 Conceptual representation of the H-filter developed in Paul Yager's lab at the University of Washington. Small solutes are filtered out of the sample stream based on their enhanced diffusivity transverse to the flow. Reprinted by permission from Macmillan Publishers Ltd: Nature [21], © 2006.

1.3.4 Surface forces and droplets Another important difference between fluid motion at the macroscale and the microscale is the enhanced significance of surface tension. The Bond number (Bo), defined as

Bo ≡

Δρ ⋅ a ⋅ L2

(1.7)

γ 12

represents the dimensionless ratio of body forces to surface tension forces at a fluid-fluid interface. Here Δρ is the density difference across the interface, a is the acceleration associated with the body force, which in most cases is gravity, L is the pertinent length scale (typically the radius of a droplet or diameter of a channel), and γ12 is the surface tension between the two fluid phases. The magnitude of the body forces relative to the surface tension forces rapidly decreases as the length scale is reduced, thereby reducing the Bond number. Fluidic control systems based on surface tension forces most commonly use changes in channel geometry or surface hydrophobicity to induce or inhibit pressure-driven flow. Surface tension forces can help drive fluid flow by exploiting capillary action in a phenomenon often described as wicking. In wicking, intermolecular forces between the fluid and a narrow channel’s surface trump those


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within the fluid such that it is drawn through the channel. A simple application of this phenomenon is that of a blood viscometer, in which the sample’s viscosity is measured based on the distance it is drawn through a narrow channel [22]. Similarly, flow can be inhibited by altering the hydrophobicity of a channel surface. Several chemical and physical processes have been used to create hydrophobic stops at desired locations within microfluidic networks, such that the flow of an aqueous solution will be impeded upon reaching that location [23]. Surface tension is particularly significant in droplet-based - also referred to as digital - microfluidic systems due to the increased surface area-tovolume ratio that accompanies decreased fluid volumes. These digital systems most often utilize two immiscible phases with the continuous phase being a hydrophobic fluid, such as mineral oil or air, and the disperse phase being an aqueous solution, as is characteristic of biological samples. Droplets help suppress unwanted Taylor dispersion and evaporative effects, and have been used for many applications, including sample transport, mixing, and particle synthesis [24]. Electrowetting is a technique in which electric potentials are used to alter the local surface wettability of a substrate at opposing ends of a droplet, thereby inducing a pressure gradient within the liquid and subsequent droplet motion. Thermocapillary pumping is another example of droplet transport exploiting surface tension, in which one end of a droplet is heated to create a surface tension gradient and induced motion [25]. Table 1.2 Important dimensionless numbers in microfluidic systems Dimensionless Definition Significance Microfluidic advantage number Reynolds Ratio of inertial Typically Re is small: ud Re = Results in laminar flow for number forces to ν most microfluidic applica(Re) viscous forces tions. Sherwood number (Sh) Bond number (Bo)

Sh =

Bo ≡

kd D

Ratio of convective to diffusive mass transfer

Δρ ⋅ a ⋅ L2 Ratio of body

γ 12

forces to surface tension forces

Typically Sh is small: Diffusion is more important with smaller dimensions. Typically Bo is small: Enables pumping via capillary pressure and droplet-based transport systems.

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1.3.5 Pumps and valves While the reduced dimensions of microfluidic systems create several advantages from a transport viewpoint, designing accurate pumping and valving control systems at this scale can be an arduous task. Nonetheless a wide variety of pumping methods for generating pressure-driven flows in microfluidic systems have been developed. The most straightforward method is direct application of externally controlled liquid sources such as syringe pumps, which are most suitable for systems containing high hydraulic resistances. Direct application of external pressure sources has also been demonstrated using both temporal and mechanical attenuation mechanisms to generate pressure differences appropriate for precise control of microscale flows. Several microfabricated pumping techniques have been developed with either off- or on-chip actuation. Serial deflection of PDMS membrane valves using external pneumatic control has gained widespread acceptance for generating pulsatile flows [26]. Similar PDMS valve designs substituting pneumatic actuation with external mechanical actuators, such as Braille pins, have also been reported [27]. Integrated actuation mechanisms, including electrostatics, piezoelectrics, electromagnetics, and thermal expansion, have been successfully implemented for actuation of a variety of other diaphragm materials. In addition to reciprocating pumps, microfabricated flow control components employing principles as diverse as thermal transpiration [28] and the Venturi effect have also been demonstrated [29].

1.3.6 Electrokinetics Electrokinetics encompasses a range of techniques for inducing motion of charged particles or conductive media by application of electric fields. Electroosmosis is the process by which the bulk fluid in a microchannel or porous media is “pumped” under application of an electric field. When an electrolyte solution is placed adjacent to a charged surface, such as glass, a layer of counter ions called the Stern layer adsorbs to the surface. Between the Stern layer and the bulk liquid is a region of intermediate charge density known as the diffuse electrical double layer. Movement of the double layer, and subsequently the bulk liquid, can be induced by electrostatic forces resulting from application of a DC potential along the channel axis. Electroosmotic flow results in a flat, plug-like velocity profile across the channel width, unlike the parabolic flow profile typical of pressure driven flow (Fig. 1.7).


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Fig. 1.7 Physical representation of electroosmotic flow. Counterions (+) in bulk solution amass near the negatively charged surface in the electrical double layer. These ions migrate with the application of an electric field, and drag the remaining bulk solution via viscous forces. Electroosmosis results in a “flat” velocity profile, as opposed to the parabolic profile of pressure driven flow.

Electrophoresis, on the other hand, is simply motion of a charged particle in solution under an applied electric field. Most biological molecules and particles carry a surface charge that will induce a force in the direction of the electric field. The electrophoretic velocity, u, of a particle is governed by u = μE, where μ is the electrophoretic mobility of the particle (dependent primarily on its size and net charge) and E is the electric field strength. Electrophoresis allows for separation of different species (e.g. DNA, proteins, etc.) based on unique electrophoretic mobilities. Several electrophoretic techniques have been developed and implemented in microscale formats for enhanced separation and enrichment of charged analytes. For example in isoelectric focusing (IEF) [30-32], species migrate through an imposed pH gradient and drift toward their respective isoelectric point the pH at which the molecule’s net charge is zero. An analogous technique is temperature gradient focusing (TGF) [33, 34], in which species are separated by their temperature-dependent mobilities along an imposed temperature gradient. Another example includes isotachophoresis (ITP) [35], in which species are separated based on their different electrophoretic mobilities as they migrate between trailing (slow) and leading (fast) electrolytes. Droplets, as well as particles, suspended in a liquid medium can alternatively be manipulated using dielectrophoresis (DEP). The principle of DEP does not rely on the charge of particle but rather the relative polarizability of the particle with respect to the surrounding medium. In the presence of a non-uniform electric field, particles possessing a greater po-

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larizability than the surrounding medium experience a force toward regions of higher field strength (positive DEP); conversely, particles less polarizable than the medium experience a force in the direction of lower field strength (negative DEP). Because the dielectrophoretic force is proportional to the gradient instead of the polarity of the electric field, both DC and AC fields can be used. AC and DC dielectrophoresis have been extensively used to trap bacteria and cells [36, 37].

1.3.7 Thermal management As is the case in most biological systems and assays, many of the processes performed on microfluidic devices are highly temperature sensitive. For example, PCR requires cyclical temperature fluctuations of 94°C, 54°C and 72°C, while most restriction endonucleases in restriction digest reactions are inactivated at temperatures above 65°C [38]. When inducing thermal fluctuations with microfabricated heating elements, it is important to ensure that the heating is in an isolated region and not adversely affecting other processes in the device. Additionally, many microfluidic devices also contain electronic components which can produce Joule heating, and therefore need to be isolated in a similar manner. Many thermal isolation techniques have been developed. One of the most common techniques is to use thermally resistant silicon barriers, such as cantilevers and bridges, to protect sensitive portions of the device [39-44]. Other materials with low thermal conductivity have been used, including quartz [45], ceramics [46] and porous silicon [47, 48]. Additional thermal isolation techniques include thermal conduits, silicon back dicing and silicon back etching [38]. Thermal conduits work by enhancing heat conduction at places where low temperatures are desired. Back-dicing and back etching involve physically isolating the heat generation regions from regions that are desired to be kept cool. Another way to thermally isolate certain regions is by selective cooling, a process in which microchannels are backed with heat sinks [49].

1.4 Device Fabrication The past few decades have generated a wealth of new materials and techniques for fabricating microfluidic devices. The myriad fabrication tools available to researchers today allows for simple processing, improved functionality, and rapid and inexpensive prototyping. Here we outline the


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standard materials and processes used in constructing microfluidic devices; these concepts will be explored in further detail in a subsequent chapter.

1.4.1 Materials Early microfluidic systems predominantly used silicon as a substrate due to the wealth of existing fabrication techniques in microelectronic production. Researchers used standard photolithography and etching processes to construct microfluidic channels of precise dimensions on silicon wafers. As research progressed, however, the focus shifted from silicon to glass substrates. Glass presented a number of advantages over silicon. Perhaps its most obvious advantage is that glass is transparent, thereby allowing visualization of on-chip processes as well as simple detection for separation assays. Additionally, glass is more compatible with electroosmotic flow than conductive silicon, which was arguably the dominant pumping mechanism at the time. From a fabrication standpoint, glass can be bonded to a second substrate more easily than silicon. In general, although the cost of glass and silicon wafers is about the same, glass became regarded as a simpler and more universal substrate for microfluidics. For many applications, glass and silicon soon gave way to polydimethylsiloxane (PDMS), an elastomeric polymer used in everything from contact lenses to bathroom caulking. When cured, PDMS behaves like an elastic solid that maintains its molded structure. The use of PDMS for microfluidic applications was first presented by George M. Whitesides in the mid1990’s [50, 51]. In this method the elastomeric monomer is poured over a master mold structure (typically a silicon wafer with an inverse photoresist structure of the channel geometry) and then cured (Fig. 1.8). The cured structures can simply be peeled from the master and bonded to a planar substrate, forming the microchannel structures. The master mold can be reused multiple times to replicate devices. Thus, this method is especially desirable due to its low cost, flexibility, and rapid prototyping. Other polymers have also been used for microfluidic applications, including polymethylmethacrylate (PMMA) and polycarbonate (PC), and are typically fabricated using hot embossing and injection molding methods. PDMS, however, remains the dominant polymer used in microfluidic fabrication. Although PDMS gained widespread acceptance in the field, glass and silicon certainly still have utility in microfluidics. Polymers typically have inferior chemical resistance, aging, mechanical, thermal, and optical properties than glass and silicon. For instance, PDMS can restrict detection of

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low wavelength fluorescence (near 400 nm). Thus for laser-induced fluorescence (LIF) detection, sensitivity is lower in PDMS than in comparative glass devices. Silicon has the advantage that it can combine on-chip electronics (i.e. via CMOS) with microfluidic networks. Also, silicon fabrication still supports the most precise geometrical tolerances, which will be increasingly important as channel dimensions continue to decrease. There is little doubt that we will see continued application of all three substrate materials (silicon, glass, and polymers) as microfluidic research proceeds.

Fig. 1.8 Schematic representation of PDMS casting using a silicon master mold with features fabricated using photoresist. Reprinted by permission from Macmillan Publishers Ltd: Nature [52], © 2007.

1.4.2 Fabrication and assembly Adopting the fabrication technology used in the microelectronics industry, microfluidic researchers frequently use lithographic techniques for creating the features needed in their device. Photolithography involves exposing a substrate coated with a photo-sensitive material, called a photoresist, to light such that the selectively developed regions can be shielded from (or subjected to) subsequent fabrication processes such as etching or deposition. Exposure transfers the 2-D pattern of desired features via a pho-


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tomask placed between the substrate and light source. Material can then be removed from the substrate using etching techniques (wet or dry). Likewise certain materials (i.e. polymers and metals) can be embossed to the substrate surface using various deposition methods. Microfluidic channels are then typically enclosed by bonding the fabricated substrate to a planar matting material. This is usually accomplished using anodic bonding (glass-silicon), fusion bonding (silicon-silicon), or by depositing a thin film of polymer or adhesive between the substrates that will bond upon hardening. More traditional fabrication approaches have also been explored for constructing microfluidic devices. Conventional machining methods (i.e. milling, drilling, cutting, and turning) that remove material from metals and hard plastics have been used to create simple microstructures, primarily for molding applications [53, 54]. The benefit of these methods is that the mold can be constructed from robust materials like stainless steel that have long lifetimes. This method is generally only suitable for simple planar channel structures with dimensional tolerances on the order of tens of microns. Injection molding is by far the most popular method used in the production of polymer parts on the order of millimeters to centimeters or larger, and has been used to produce microfluidic devices [55, 56]. A polymer melt is injected under high pressure into an evacuated die cavity containing the desired master mold, and subsequently cooled and solidified such that it can be separated from the mold. While probably not practical for simple prototyping, injection molding would be a desirable fabrication tool for high throughput production of commercial devices. Threedimensional polymer microdevices have been fabricated using stereolithography, a bottom up fabrication process that uses a focused laser to optically cure polymer at precise coordinates in real time [57]. The advantage is that the fabrication is very simple in comparison to other techniques as it requires no masks; the structure is defined using a CAD program which allows for rapid and inexpensive prototype adjustments. However, this system is somewhat advanced and might be time consuming and cost prohibitive for the construction of simple microfluidic devices.

1.5 Biological Applications While microfluidics seemingly has application in many areas, such as manufacturing and aerospace disciplines, thus far it has almost exclusively been pursued in the biological sciences. Here we briefly explore some of

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the most prevalent biological applications as a prelude to the more in-depth reviews presented in subsequent chapters of this text.

1.5.1 Genetic analysis (DNA/RNA) In the past few decades, genomic research has yielded powerful techniques with applications ranging from genotyping to clinical diagnosis. Completion of the human genome sequencing brought with it a high demand for rapid, high throughput genetic analysis. Microfabricated systems have become desirable solutions for rapid and inexpensive genetic assays that offer low sample consumption, low fabrication cost, and short operating time. Expanding on early developments in modular reaction and separation systems, integrated genetic assays have evolved into fully-functional microfluidic platforms. As the key component of most genetic assays, PCR has been studied extensively in microsystems. PCR microsystems can be categorized into two major groups: static flow-based systems, in which the reaction mixture remains in a chamber and is heated with external or on-chip heaters, and continuous flow-based systems, in which the reaction mixture flows through different temperature zones at a controlled flow rate [58]. Novel approaches, like thermal convection-based PCR, have also been reported [59]. Other genetic analysis techniques, including DNA sequencing, reverse transcript PCR, restriction digestion, ligase detection reaction (LDR), and capillary electrophoresis (CE) have also been successfully performed in microdevices. Recently, integrated microsystems that contain sample preparation, fluid handling, bioreaction, and product detection (either electrophoresis or microarray hybridization) have been constructed, providing the capability to quickly collect genetic information from raw biological samples [60]. In addition to genetic analysis, microfluidic devices combined with microelectronics have also been applied to investigate the dynamic behavior of single DNA or RNA molecules, and the interaction between DNA/RNA and proteins.

1.5.2 Proteomics In the post-genomic era, proteomics has become one of the most studied and challenging areas of biological interest. A map of the human proteome, similar to that of the genome, would provide enormous knowledge to biologists studying the effects of the environment, diseases, and drugs on the body [61-63]. Such a goal requires the development of rapid and


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high throughput devices for molecular investigations of cellular compositions. Proteins are also of interest for clinical diagnostics, in which relative concentrations of protein biomarkers in physiological fluids can signal the presence of a systemic disease [64, 65]. Due to the diversity of amino acid chains in their composition, proteins are more complicated and show a much wider range of structures than DNA or RNA. Also unlike DNA and RNA, proteins have no self-amplification procedure; therefore dilute samples must be preconcentrated or enriched prior to analysis in order to overcome instrumental limits of detection. Moreover, proteins are highly unstable thermally and physically meaning careful design and operation is needed when manipulating proteins in microfluidic devices. Microfluidic protein analyses require sample pretreatment steps (i.e. extraction, dialysis, and enrichment) followed by the assay itself (i.e. immunoassays, enzymatic assays, or electrokinetic focusing and separations). While a great deal of research has been conducted on individual phases of proteomic analysis, full-integration is an arduous task and remains the objective of several efforts. With integration in mind, interfacing microfluidic devices with traditional identification methods may prove particularly advantageous for proteomic research. For example, mass spectrometry methods, such as MALDI-TOF and ESI-MS, are used frequently for characterizing unknown protein samples. Therefore to facilitate rapid mass analysis of proteins, many researchers are developing devices for interfacing microfluidics with mass spectrometers [66].

1.5.3 Cellular assays Due to the controllability and reproducibility of microfluidic systems, miniaturization of cell cultures and assays has been studied intensively. Microfluidics has been shown to provide more genuine in vitro environments than traditional cell culture techniques due to efficient heat and mass transport. Serial processing and parallelization have also enabled high throughput assays on a single chip [67]. For eukaryotic cells, biologically compatible materials (such as PDMS) can be structured as an extracellular matrix using the microfabrication methods mentioned earlier. This control of surface shape has been used to investigate cell physiology in various cultivation environments [68]. In addition, taking advantage of well-controlled laminar flow enables precise microfluidic cell treatment, such as chemical gradient or temperature steps.

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These subcellular changes of cultivation environment are generally driven by diffusive mixing in a laminar flow [69, 70]. The combination of microfluidics and cell cultivation is also widely used in the field of microbial research. For example, a microfluidic chemostat for monitoring microbial growth was developed by S.R. Quake and his colleagues [71]. Bacteria immobilization, control of cell morphology, and dynamic bacterial population analyses have also been facilitated with microfabricated devices. Another important research area is that of single cell assays. Single cell assays eliminate the ensemble averaging effect of cell populations and enable the precise investigation of individual cells. A wide range of isolation methods have been developed, such as hydrodynamic focusing, microdroplet generation, microwell arrays, and physical barrier arrays [72]. Cell sorting and screening is also one of the most rigorously investigated topics. Cells can be screened and sorted from a continuous flow according to electrophoretic mobility, refractive index, or size. In particular, microscale flow assisted cell sorting (micro-FACS) systems implementing DEP or electroosmotic flow have been extensively pursued [73-75].

1.5.4 Drug delivery and compatibility In addition to their extensive applications in analysis and diagnosis, microfluidic devices also offer a host of benefits for in vivo applications, particularly for drug delivery. Microfluidic drug delivery devices not only deliver proper concentrations of functional drugs to certain target sites with controllable release rates and dosage, but also present some other advantages: reduced size and power consumption, simple operation, and the ability to achieve complex release patterns (e.g. continuous or pulsatile release). Drug delivery microsystems can be categorized into three main groups [76]: (a) biocapsules and microparticles for controlled and/or site-specific drug release, (b) microneedles for transdermal and intravenous delivery, and (c) implantable microsystems. Biocapsules usually contain micro/nano-porous biocompatible membranes for drug encapsulation. Today, microparticles with various size and shape can be produced using standard microfabrication techniques. Combined with proper coatings, these microparticles provide effective site-specific delivery, especially for peptide and protein-based drugs. Microneedles have been fabricated as both 1-D


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in-plane arrays and 2-D out-of-plane arrays, providing transdermal drug delivery with minimal tissue damage and negligible pain sensation [77]. Most of the microneedle-based devices are also integrated with certain micropumping components to control the drug dosing. Implantable microsystems are more suitable for therapies that require many injections for a certain period of time, because they can not only reduce the injection times, but also precisely control the rate of drug release. With the development of implantable microsensors [78] comes the potential to build “smart” drug delivery device (Fig. 1.9). These devices will function without any human intervention by: (a) real-time monitoring of target physiological conditions in a patient’s body using microsensors, (b) converting these conditions into some detectable signal, and (c) controlling the drug release through microactuators based on the signal analysis [76]. The resultant closed-loop drug delivery can provide the patient with a selfregulated treatment regiment. One important issue that has to be considered in designing microsystems for biological applications is biocompatibility of the device substrates how to interface these man-made devices with the relevant biological in vitro or in vivo environments so as to ensure proper functionality for a desired period of time. Devices must be able to withstand attacks from bioreactions and/or bodily immune responses (for in vivo applications), and should not cause any unwanted reactions or inhibitions. There is no standard list of biocompatible materials due to the differences in specific biological applications, as well as other details such as fabrication processes and the implanted location and lifetime of the device. Biocompatibility is a difficult property to control (and define) and thus remains the subject of much research.

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c

Fig. 1.9 Implantable drug release device in which the gold membrane anode covering in (a) is removed in (b) via the application of an electric field to initiate release from a reservoir. The plot in (c) shows the intermittent release rate of the device over several days. Reprinted by permission from Macmillan Publishers Ltd: Nature [79], © 1999.

1.6 The Future 1.6.1 Potential demand/market for microfluidic devices The microfluidic technological impact has long been anticipated, but use of lab-on-a-chip devices is still limited. Although a few LOC products are now commercially available for applications such as DNA analysis, pro-


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tein crystallization, and performing simple chemical reactions, the field continues to search for a so-called “killer application” [80]. For healthcare alone, the potential market for simple and inexpensive diagnostics seems to be incredibly promising. In the future, healthcare providers will likely shift from treating diseases to anticipating and preventing them. Widespread screening or testing will be a necessity for such a system, and microfluidic platforms are the most plausible technology for successful implementation. Consider, for example, genetic screening of newborn children for indications of potential health risks. For a quick, orderof-magnitude approximation of the number of tests that might be required each year for such analysis in the United States alone, we can multiply the approximate number of genes times an estimated number of variants per gene [81], times the number of births each year [82] and arrive at 103 x 102 x 106 = 1011 tests per year. Of course this number represents a broad approximation, and the term “test” is not equivalent to “device” (i.e., one device can perform many tests), but the number does imply that there is potentially a large market for diagnostics that are as simple to acquire and operate as commercial home pregnancy tests.

1.6.2 Current products We need to remember that the field of microfluidics is still very much in its early adolescence. However, a number of companies are now developing and marketing microfluidic lab-on-a-chip systems for use in various areas such as biomedical research, environmental testing, and medical diagnostics. Micronics mass-produces disposable microfluidic ‘lab cards’ that can be used in the development of devices for various applications, most notably low-cost disposable cards to analyze disease in the field. Another established microfluidics-based company is Agilent, whose 2100 Bioanalyzer® is a successful device for analyzing DNA, RNA, proteins and cells using a microfluidic format. Caliper Life Sciences produces the LabChipTM system, which has become widely used by pharmaceutical firms for cellular and enzymatic screening. Other successful microfluidic products used in research labs and hospitals include the GeneXpert® system by Cepheid for amplifying and detecting target DNA, the Triage® BNP test by Biosite for assessing the severity of a heart failure, and the iSTAT® 1 system by Abbott for conducting rapid, comprehensive blood screening with minimal sample volumes. Microdevices for liquid dispensing have been developed in various formats by TTP Labtech, Perkin Elmer, Tecan, and Labcyte. These liquid dispensers can manipulate vol-

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umes as low as 5 nl with high precision. Silex Microsystems in Sweden has been adopting MEMS-based techniques to produce microfluidic labon-a-chip devices for the life sciences market. HandyLab, based in Ann Arbor, MI, also manufactures devices for nucleic acid testing. Fluidigm produces lab cards for protein crystallization and multiplexed genotyping using their multilayer soft lithography technique (Fig. 1.10). Nanogen offers a microfluidics-based array platform for DNA hybridization research to help users selectively probe target DNA. Unlike companies that commercialize microsystems for specific applications, Micronit Microfluidics is a commercial supplier and fabricator of microfluidic devices for researchers worldwide. The Stanford Microfluidics Foundry and the Michigan Nanofabrication Facility are other microfluidic fabricators known for soft lithography. These commercial foundries produce customized microfluidic devices from various materials, tailored to customers’ demands.

Fig. 1.10 Fluidigm’s BiomarkTM 48.48 Dynamic Array device for multiplexed genotyping. Reproduced with permission from www.fluidigm.com.

1.6.3 Challenges and the future Existing microfluidics-based companies have opened the door to device commercialization, but it seems that they have just scratched the surface of the potential market. So, just what is holding the field back? Why haven’t we seen an explosive flux of new products into the market? The answer, as it is with most emerging technologies, appears to be cost. Microfluidic devices are relatively expensive to manufacture due to the equipment and processes typically required. Additionally, these devices are competing with conventional platforms with which most customers are accustomed.


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Microfluidic technology must significantly outperform or cost less than current products in order for customers to justify the switch from conventional technology. The market will forever be “user-driven” rather than “technology-driven”. User-friendliness is another major hurdle to microfluidic device commercialization. Microfluidic systems must be easily operated by non-experts. User-friendliness involves many challenges such as system integration, interfacing, and packaging. While these issues remain hurdles toward commercialization, they still receive little research or funding support. Devices must have macroscopic inputs and should employ user-friendly diagnostic concepts such as the simple yes/no indicators for the presence of antigens, antibodies, viruses, or other biological targets used in commercial products (e.g., pregnancy tests). Running real-life fluid samples like blood and saliva through a microfluidic device, however, typically brings more problems than using a purified laboratory solution. Therefore researchers also need to design devices such that they are operable in real-world environments, rather than solely in the laboratory. The future of microfluidics holds enormous potential as researchers continue to bring the technology out of the laboratory and into our real life. The innate advantages of microfluidics are too hard to ignore; we will undoubtedly see microfluidic devices replace conventional techniques as the field is further explored. Commercial success will drive future research efforts, and the field will expand with influences not only in the biological arena, but other disciplines as well. The remainder of this text will further explore many exciting and diverse research areas associated with this budding technology.

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Chapter 2 Materials and Microfabrication Processes for Microfluidic Devices

Arash Noori, Sarvesh Upadhyaya, and P. Ravi Selvanganapathy McMaster University, Hamilton ON L8S 4L7, CANADA Correspondence should be addressed to: P. Ravi Selvanganapathy ([email protected])

Keywords: Microfluidics, microfabrication, Silicon, Glass, Parylene, PDMS, Hydrogel, Paraffin, Polyimide, bulk micromachining, surface micromachining, micro injection molding, hot embossing, micro stereolithography

Abstract This chapter elaborates on the varied materials and microfabrication techniques used in the manufacture of microfluidic devices. The origins of these techniques traces back to semiconductor and precision machining industries. This chapter details microfabrication processes such as bulk and surface micromachining for silicon, glass and polymeric materials. It also discusses precision machining techniques such as micro injection molding, hot embossing, micro stereolithography. Over the past few decades, these techniques have been modified to suit the complexity and precision required in the 3-D construction of microfluidic devices. New materials such as parylene, poly-dimethylsiloxane, paraffin and hydrogels have been in-

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Noori, Upadhyaya, and Selvaganapathy

troduced and microfabrication techniques developed that enhances the functionality of microfluidic devices. This chapter provides a comprehensive overview of material properties, microfabrication procedures and examples of its application in microfluidic devices.

2.1 Introduction Microfabrication technology originated in the semiconductor industry in response to the need to manufacture integrated circuits. These techniques were primarily lithography based and were developed for silicon based materials [1]. Since integrated circuits have their critical elements (transistors) in microscale dimensions and are packed tightly together, the fabrication technologies developed were targeted at achieving precision in the sub micrometer scale. Simultaneously, there has been a significant effort in the precision machining industry to develop materials and processes that achieve microscale precision for applications in a number of areas including micromotors, sensors, micro-optics and other precision components. A number of materials including metals, polymers and ceramics and process such as electrodischarge machining, precision milling, single point diamond turning, and laser micromachining have been developed over the years [2]. Several of these methods and processes for manufacture have been adopted and combined over the past 15 years to create microfluidic structures for applications in areas as diverse as medical diagnostics, drug delivery, drug discovery, analytical chemistry, combinatorial synthesis, molecular diagnosis, and specialty chemical manufacturing [3]. Laboratory based analytical methods used in areas of medical diagnostics and drug discovery are the gold standard in accuracy but are slow, have low throughput, and have high cost per analysis. Similarly, yields in special chemical manufacture are low due to the lack of micro level control over unit operations and residence times. Microfluidic devices offer the promise of miniaturizing and integrating multiple laboratory unit operations within a single chip. Miniaturized Lab-on-Chip (LoC) devices consume small volumes of samples and expensive reagents and can be made portable and available at the point of care. They also reduce the time of analysis significantly as unit operations like heating, mixing, and metering can be performed more accurately and quickly in smaller volumes thus providing real-time analysis. In the case of microreactors, they allow precise definition of process parameters such as interaction time between reactants by the structural design of the reactor, enabling high yields and fea-

Materials and Microfabrication Processes

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sibility of certain reactions that were not possible before. Using microfabrication, several devices can be produced in parallel with the same processing steps, reducing their unit cost. Furthermore, the structural features and hence the functioning of the devices are accurate and repeatable which leads to accurate analysis and higher degree of confidence [4]. Several materials have been used in the construction of microfabricated devices such as silicon, glass, polymers, metals and ceramics. The most widely used material has been silicon, primarily due to the fact that early researchers were more familiar with the material and the processes involved in structuring it to the necessary shape. Glass is also used extensively in microfluidic devices, especially those targeted for biological applications, as most of the biochemical reactions have been characterized in glass. Lately, polymers have been incorporated into microfluidic devices due to the variety of surface properties obtainable and their ease of fabrication. This chapter deals with some of the widely used materials in microfluidic applications along with their processing techniques.

2.2 Silicon Based Materials The first miniature microfluidic system was a gas chromatographic system, microfabricated in silicon substrate, which was able to separate and analyze simple gas mixtures [5]. The system consisted of a sample injection valve and a 1.5m long separating capillary column fabricated on a silicon substrate and a nickel film as the sensing element of the detector. Silicon was the preferred material for initial microfluidic systems due to the availability of fabrication techniques and equipment from the microelectronics industry. Around the same time, research was ongoing at IBM for the development of inkjet printer heads [6, 7, 8] in silicon using anisotropic etching techniques. These developments can be seen as the origins of microfluidics. Over the years, with the development of more sophisticated fabrication techniques, a number of these microfluidic components have been integrated on a single substrate to automate and perform a complex series of unit operations for biochemical analysis [9]. Applications include DNA analysis [10], gas chromatography [11], chemical microreactors [12], forensic analysis [13], environmental analysis [14], and drug discovery [3]. Micro-total-analysis-systems offer a number of potential advantages when compared to conventional systems, including lower fabrication costs due to massively parallel fabrication techniques, reduced reagent consumption and dead-volume, and improved performance due to the possibility of per-

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forming multiple analyses on a single chip [15]. Although other materials such as polymers are gaining in popularity, silicon continues to play a crucial role in the fabrication of microfluidic devices. A number of companies such as MEMSCAP, Colibrys, Intellisense, Integrated Sensing Systems Inc. (ISSYS), Innovative Micro Technology (IMT), Micralyne Inc., Micronit BV, Silex Microsystems and GE sensing offer foundry services or produce limited volumes of several microfluidic devices in silicon. Application areas include microneedles for drug delivery; devices for cell sorting, filtration and purification; DNA sensing and amplification devices; and sensing devices for flow, pressure, temperature, pH, and humidity. Some of the more commonly used materials are silicon, silicon-di-oxide, and silicon-nitride. Their material properties are shown in Table 2.1. Table 2.1 Material. properties of commonly used silicon based materials in microfluidics Properties Electrical Dielectric strength Resistivity Dielectric Constant

Silicon

Silicon Oxide

Silicon Nitride

1000ohm-cm 11.8

107 V/m 1016 ohm-m 3.9-4.3

107 V/m 1014 ohm-cm 7.5

Mechanical Tensile strength Young’s modulus Poisson’s ratio Elongation to break Density Hardness

2 GPa 160 GPa 0.27 1.2% 2.33 g/cm3 11.9-13 GPa

75 GPa 0.17 2.19 g/cm3 9 GPa vickers

260-330 GPa 0.23 - 0.27 3.1 g/cm3 16-18 GPa vickers

Thermal Thermal Conductivity Melting Temperature Coefficient of thermal expansion Specific Heat Capacity

150 W/m.K 1414 C

1.4W/m.K 1610 C

25-36 W/m.K 1900 C

2.49 x10-6 /C 0.702 J/g.K

0.5 x10-6 /C 1.0 J/g.K

2.8-3.2 x10-6 /C 0.54-0.7J/g.K

Other Properties: Index of refraction

3.42

1.46

2.05


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2.2.1 Micromachining of silicon Silicon micromachining can be classified into two categories, bulk micromachining and surface micromachining. Of these two technologies, bulk micromachining is the most widely used and most commercially successful technology [16]. It involves the selective removal of the substrate material, in the regions unprotected by a masking layer, which allows the definition of features in the substrate. The masking layer itself is patterned using photolithographic process. Bulk micromachining has found applications in the manufacture of various devices such as pressure sensors and ink-jet print heads [17]. Conversely, surface micromachining is used to create microstructures on top of the substrate, rather than in the substrate itself, using photolithographic process. It involves the deposition of thin films on a patterned sacrificial layer, which is then etched away, leaving the thin film as the structural material. Lately, these surface micromachined techniques have gained importance in fabrication of microfluidic networks and components. 2.2.2 Bulk micromachining Micromachining processes that etch into the silicon substrate for device fabrication are referred to as bulk micromachining techniques. Bulk micromachining of silicon combined with other processes (such as metal deposition to create electrodes, chemical mechanical polishing, and subsequent wafer bonding) is used for the fabrication of microfluidic devices. Bulk micromachining processes can be further divided into etching using liquid etchants (wet etching) and etching using gaseous etchants (dry etching) [18]. The etch profile in these processes can be isotropic or anisotropic. Isotropic etchants etch uniformly in all directions, whereas anisotropic etchants are direction dependent and etch some directions preferentially over others [19]. As a result of this, isotropic etches result in rounded cross sections, whereas anisotropic etches result vertical sidewalls or those constrained by the crystal planes of the silicon substrate. This is shown in Fig. 2.1. Etch rates are characterized and etching process carried out for the required duration of time to obtain the desired feature dimension. Choice of etchant, masking material, and etch stop layer are all critical in the design of the bulk micromachining process [18]. Structures that have been fabricated by bulk micromachining include micro channels, micro wells, fluidic interconnects, micropumps, valves, mixers and reaction chambers for microfluidic applications.

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Fig. 2.1 Cross sectional profiles of different etch methods: a.) Isotropic wet etch, b.) Anisotropic wet etch, c.) Isotropic dry etch, d.) Anisotropic dry etch.

2.2.2.1 Wet isotropic etching

Wet etching of silicon is widely used because of its fast etch times, low cost, low-complexity, and the availability of masking materials [20]. A commonly used isotropic wet etchant is a mixture of hydrofluoric (HF), nitric (HNO3), and acetic (CH3COOH) acids also known by the acronym HNA. When a silicon wafer is exposed to the etchant, it is oxidized by the nitric acid. The product is then etched by the hydrofluoric acid [19]. The details of this etch can be summarized into the following four basic steps. The process begins with the injection of electron holes into the Si to form higher oxidation state Si2+ or Si+. Hydroxyl groups (OH-) then attach to the positively charged Si, and initiate the reaction of the hydrated Si with the complexing agent in the solution. Finally, the reacted products are dissolved into the etchant solution. Therefore, any etching solution must provide a source of electron holes, hydroxyl groups, and a complexing agent whose reacted species are soluble in the etchant solution [18]. Depending on the ratio of the acids in the mixture, etch rates of silicon in HNA can vary from 0.1 to over 100 µm/min. Uniformity of etches is typically difficult to control but improves with stirring [19]. 2.2.2.2 Wet anisotropic etching

Anisotropic etchants such as potassium hydroxide (KOH), sodium hydroxide (NaOH) and Ethylenediamine - pyrocatechol (EDP) have etch rates dependent on the orientation of the crystallographic planes on the silicon wafer. On a silicon wafer, the [111] planes are at an angle of 54.74 degrees


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for [100] wafers and 90 degrees for [110] wafers. In the anisotropic etchants, the etch rate along [111] plane is generally the slowest by 2-3 orders of magnitude. This property can be exploited with careful design to produce anisotropic structures and smooth sidewalls. For instance, at 80oC a mixture of potassium hydroxide (KOH) in water and isopropyl alcohol has an etch rate of about 2.1µm/min in the [110] plane, 1.4um/min in the [100] plane, but only 0.003µm/min in the [111] plane [16]. The higher etch rates in the [100] and [110] directions result in the exposure of the slow etching [111] planes over time. Figure 2.1 shows the orientation-dependent etching of a [100] silicon wafer with a patterned silicon dioxide mask. Using anisotropic etching, it is possible to accurately predetermine the result if the characteristics of the etchant are known [21]. With a proper mask design and orientation, one can obtain perfectly aligned square or rectangular cavities and pits, V-grooves, as well as holes or channels with vertical sidewalls [18]. Some of the earliest microfluidic devices were fabricated using anisotropic etching. Bassous et. al. [7] fabricated arrays of nozzles with square orifices for ink-jet printing applications on [100] wafers using an Ethylenediamine - pyrocatechol (EDP) etching solution. The nozzles consist of truncated square pyramidal cavities bounded by the four convergent [111] planes and 2 [100] surface planes as shown in Figure. 2.2. The anisotropic etches provide a very high degree of symmetry as a result of the crystallographic perfection of the starting wafer and a well controlled etch rate. The fabrication is carried out using conventional Si-processing techniques. The wafer is chemical-mechanically polished, cleaned, and thermally oxidized to form a 0.5µm thick SiO2 film. Photoresist is applied to both surfaces and patterned. Subsequently, the square openings are defined and etched in the SiO2 film to reveal the silicon wafer. The wafers are then etched in a solution of EDP, until the openings appear on the back side of the wafers. Finally, the wafers are cleaned, the oxide is stripped and the wafers are oxidized to provide a uniform oxide coating throughout the structure.

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Fig. 2.2 Anisotropically etched pyramidical inkjet nozzle in silicon (Reproduced from [7] with permission of ECS – The Electrochemical Society).

In the previous example, the etching stopped when the orifices appeared on the back side of the wafer. Another method involves the use of timed etches, which may be used when tolerances for the final structure are not important or when the etch rate is well defined and reproducible [20]. However, when accuracy and tolerances are crucial, doping induced etch rate variations can be used as an etch stop. Highly boron doped silicon has a greatly reduced etch rate in all alkaline etchants [18]. By selective boron (p++) doping through a mask, regions can be made resistant to etching, while undoped regions will be etched away. For boron doping concentrations of ≥20x 1020cm-3, the KOH silicon etch rate of 0.7-3.0 µm is reduced by a factor of 20 [18]. This allows for accurate control of vertical dimensions, allowing the fabrication of membranes and other complex threedimensional microstructures [18]. The primary advantages of the ion implanted etch-stop techniques are independence of crystal orientation, smooth surface finish, and the ability to fabricate complex microstructures [20]. The main limitation is the maximum depth of ~15µm practically that is achievable using dopant diffusion [22]. Another method for the fabrication of microfluidic channels - developed by Chen et al [23] - incorporates doping, anisotropic etching, and isotropic etching for the fabrication of buried microchannels in silicon. The process flow is shown in Figure 2.3. The fabrication process begins with a p-type [100] silicon wafer with a patterned oxide layer that undergoes shallow boron diffusion to form a 2µm highly boron-doped layer on the surface. Using reactive ion etching (RIE), a chevron pattern is opened in the boron layer to define the channel structure. The features of the chevron structure are narrow enough to be undercut quickly in the subsequent wet etch, while preserving silicon in the bridge. To open up the channel structure, an


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anisotropic silicon etch (EDP) is used to undercut this chevron structure as shown in Figure 2.4. The sidewalls of the flow channel are [111] surfaces, which have the slowest etch rate in EDP. Therefore, the width of the flow channel is determined by the area defined by the patterned chevron bridges instead of lateral etching. Although chevron patterns are used in this case as openings, other shapes could be used as well. After the channel is etched in the silicon, deep boron diffusion is performed to define the probe shank. Sealing of the channel is accomplished using thermal oxidation of a 0.5µm of thermal oxide and LPCVD dielectrics. Figure 2.4 shows the cross section of the completed sealed channel.

Fig. 2.3 Fabrication process for chemical delivery probe using undercutting (Reproduced with permission from [23] ©[1997] IEEE)

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Fig. 2.4 Cross section of chemical delivery probe fabricated using chevron pattern and undercutting (Reproduced with permission from [23] ©[1997] IEEE).

2.2.2.3 Dry etching - Deep Reactive Ion Etching

Dry etching of silicon can be achieved using Deep Reactive Ion Etching (DRIE). In this technique ions and other chemically reactive species generated in low pressure plasma are accelerated towards the substrate producing anisotropy and enhancing reactivity. DRIE allows for the fabrication of very high-aspect-ratio structures with practical etch depths of 1mm and etch rates in the order of 2-3 µm/min [22]. A variation of DRIE, which includes alternate etching and side wall passivation steps to enhance anisotropy, was developed by Bosch and is generally referred to as the Bosch process. The process relies on an inductively coupled plasma source and an alternating etching and deposition cycle [24]. During the etch cycle, the wafer is exposed to SF6 to etch the exposed silicon. A protective polymer film consisting of CF2 molecules is then blanket deposited using a C4F8 gas source. This is followed by another etch in which the protective polymer at the bottom of the trench is removed by SF+ ions, without damaging the protective film on the sidewalls [20]. This process is repeated to obtain very high-aspect-ratio structures with anisotropies in the order of 30:1 and sidewall angles of 90+/-2 degree. Furthermore, it provides polymer and silicon dioxide selectivities of 50-100:1 and 120-200:1, respectively [22]. The DRIE process has been used extensively for the fabrication of through holes and deep microchannels in microfluidic devices. Li et al. [25] used DRIE in their fabrication of a microfluidic gas centrifuge for the separation of dilute gas mixtures. Figure 2.5 shows the fabrication process for the device. First, a 2µm layer of oxide is grown on the backside of a silicon wafer to act as an etch mask. After this, a 1.5µm layer of photoresist is deposited and patterned on the front side of the wafer, and 5µm deep gas concentrating channels are etched using reactive ion etching. A 6µm layer of photoresist is then spun and patterned for the inlet and outlet ports, after


45

which the oxide is etched with buffered HF. These unprotected openings serve as the locations where the DRIE will take place. After the DRIE is performed, the resist and oxide are stripped, and the silicon wafer is anodically bonded to a Pyrex wafer to seal the channels. As a final step, metal capillaries are inserted into the inlet and outlet ports in the silicon wafer to serve as the fluidic interfaces. DRIE was used in the fabrication of this device as existing fabrication methods of isotope separation devices relied on elaborate and costly process of stacking of photo-etched metal foils or micromachining using a combination of lithography, electroplating, and injection molding known by its German acronym LIGA.

Fig. 2.5 Fabrication process of microfluidic device using DRIE for the creation of interconnect holes. Reprinted from [25] with permission from Elsevier.

46


2.2.3 Surface micromachining Surface micromachining differs from bulk micromachining in that a deposited film acts as the structural layer, rather than the substrate itself. The process flow for surface micromachining is outlined in Figure 2.6. A sacrificial material is deposited onto a substrate and patterned. Subsequently, a film is blanket deposited onto the device to define the structural layer. After this, the sacrificial material is removed, leaving behind the desired microstructure [26]. The process requires a compatible set of structural materials, sacrificial materials, and chemical etchants with high etch selectivity such that etching of the sacrificial material does not affect the structural material [16]. It is required that the sacrificial materials have good mechanical properties to avoid problems in the fabrication process [16]. Finally, the structural material must offer the appropriate chemical and mechanical properties for the desired application.

Fig. 2.6 Surface micromachining process.


47

The polysilicon and silicon dioxide material combination is commonly used for the fabrication of microfluidic devices. The oxide is deposited onto the substrate and acts as the sacrificial material, with the deposited polysilicon acting as the structural material [26]. The deposition of polycrystalline silicon is typically achieved by gas-phase decomposition of silane using low-pressure chemical vapor deposition at temperatures in the range of 585 - 625oC. The wafer is washed using an HF solution in which the oxide is dissolved without affecting the polysilicon structure [26]. Conversely, silicon nitride can also be used as the structural layer with phosphosilicate glass (PSG) and silicon dioxide acting as the sacrificial material [27]. A combination of surface micromachining, bulk micromachining, and doping technologies has been used for the development of silicon processed microneedles by Lin et al. [28]. The process begins with a lightly doped ptype [100] oriented wafer which is selectively doped to make a 12 µmdeep heavily boron doped p-type region using a thermally grown oxide that is patterned to act as a masking layer. When the doping is completed, the oxide mask is stripped and a 400-nm-thick layer of SiO2 is thermally grown and a 600nm layer of nitride is deposited for passivation of the surface. A 600nm thick phosphorus-doped polycrystalline silicon layer is then deposited, patterned and etched, leaving behind the polysilicon resistors. After this, the polysilicon layer at the back side of the wafer is etched away and a 150nm protective layer of nitride is deposited to cover the polysilicon resistors during EDP etching. These steps prepare the substrate for bulk micromachining at the end of the process flow. Following these steps, the surface micromachining fabrication of the microchannel is performed. First, a 5µm thick layer of PSG is deposited, followed by a 3µm layer of Low temperature oxide (LTO), which improves adhesion to the photoresist and shields the interface from HF. Using a photoresist, which is subsequently stripped, the microchannel is patterned etched in buffered HF, leaving the cross section shown in Figure 2.7 (b). To create etch openings for the HF to remove the sacrificial PSG and LTO inside the microchannel, a 1µm layer of LTO is deposited to allow patterning of the openings. A 1µm layer of nitride is then deposited and patterned to create the etch holes. These holes are etched in a plasma etcher, providing the cross section as shown in Figure 2.7 (c). The etch channels are sealed by the deposition of a 1.5µm thick layer nitride after the microchannel has been fully cleared. In order to enable the separation of the needle, etch windows are opened using a plasma etcher which stops at the 400-nm-thick oxide layer. This is shown in Figure 2.7 (d).

48


(a)

(b)

(c)

(d)

(e)

Fig. 2.7 Process flow for fabrication of silicon processed microneedle (Reproduced with permission from [28] © [1999] IEEE).

Subsequently, the back side of the wafer is now patterned, which opens up the etching areas to free the microneedles. A timed EDP etch is used to reduce the silicon wafer thickness to 120µm, after which the wafer is immersed in a buffered HF solution which etches the opened SiO2 areas as shown in Figure 2.7 (e). Again the wafer is etched in EDP to reduce the 120µm thickness down to 50µm at the shank end of the microneedle. This fully releases the needle and completes the fabrication process, with the final results show in Figure 2.8.


49

Fig. 2.8 Silicon processed microneedle (Reproduced with permission from [28] © [1999] IEEE).

2.3 Glass Based Materials Glass is the most suited material for biological applications, primarily because of its favorable material properties (chemically inert, excellent insulator, optically transparent, and low auto-fluorescence), its close association with the life sciences, and the availability of virtually an infinite variety of glasses. Table 2.2 provides some of the important material properties of glasses mainly used in microfluidics. However, microfabrication techniques for glass have not advanced as much as those for silicon and it is difficult to produce high aspect ratio anisotropic structures in glass due to its amorphous nature. Early examples of microchannels constructed in glass were used as electron multipliers in X-ray applications [29] and as micropipettes for patch clamp applications [30]. Photosensitive glass has also been used in these applications to form high aspect ratio microchannels through UV exposure through a mask [31]. These methods involve heating glass capillaries and applying tensile force to draw a fine filament in micrometer dimensions and hence are not amenable to the parallel fabrication that is typical of photolithography. Subsequently, in the 1980s, unstructured thin glass was used as an actuating membrane in several silicon micropumps [32]. Actual structuring of glass using lithography for creating a network of microchannels, which is typical of microfluidics these days, began with its application in electrophoresis chips in the early 1990s [33]. Since glass capillaries were used widely in capillary electrophoresis and all the protocols had been standardized on them, early microfluidic systems were made of glass to replicate the surface characteristics and hence performance of macrosystems. Subsequent to this, development of a number of microfluidic devices

50


for diverse applications such as flow cytometry [34], micellar electrokinetic capillary chromatography [35], protein pre-concentration [36], electrospray coupling for mass spectrometry [37], and integrated systems for amplification, separation and detection of genetic materials [38] were made using glass as the structural material. A number of companies such as Micralyne Inc., Micronit BV, Innovative Micro Technology (IMT), LioniX BV, offer foundry services or produce limited volume of several microfluidic devices in glass. Table 2.2 Material properties of commonly used glass based material in microfluidics Properties

Soda lime

Electrical Dielectric strength 10 (MV/m)

Quartz

Pyrex Foturan (Corning 7740)

8

14

1.2

Resistivity (ohm-cm)

4x106

7.5x1015

8x1010

8.1x1012

Dielectric Constant

7.3

3.75

4.6

6.5

20

48

20.7

60

64

78

0.20

0.22

2.23

2.37

Mechanical Tensile strength (MPA)

Young’s modulus 70 (GPa) Poisson’s ratio

0.25

86.7 – 107.2 (direction dependent) 0.17

Density (g/cm3)

2.5

2.2


51

Table 2.2 (cont.) Properties

Soda lime

Thermal Thermal Conduc- 1.12 tivity (W/m°C)

Quartz

Pyrex Foturan (Corning 7740)

1.4

1.1

Melting Tempera- 700°C (soften- 1750 °C ture ing) 9.2 x10-6 /C Coefficient of thermal expansion Specific heat 0.21 (J/g.K) Other Properties Index of refraction 1.52

1.35

0.54 x10-6 /C

820°C (soften- 465°C (transing) formation temp.) 3.3 x10-6 /C 8.6 x10-6/ C

0.72

0.75

0.88

1.47

1.47

1.515

2.3.1 Microfabrication in glass Microstructuring of glass is mainly accomplished by bulk micromachining using isotropic wet etching. This process produces an isotropic etch, the aspect ratio of which can be controlled by the amount of stirring present in the etching chamber. This creates the pattern of the microfluidic network on the glass substrate. Fluidic access holes at reservoirs are created by a number of methods including, laser micromachining, ultrasonic drilling, abrasive powder machining, and drilling. Subsequently, the processed wafer is bonded with another glass wafer through anodic or fusion bonding to form a closed microfluidic network. Functional materials such as electrophoresis gels, porous polymers for chromatography, environmental sensitive hydrogels, and phase change actuation materials are incorporated in the microfabricated channels individually in post-processing. 2.3.1.1 Wet isotropic etching

Glass is generally indestructible to chemical attack from a wide range of chemical agents. However, a few chemicals such as hydrofluoric acid, concentrated phosphoric acid (when hot, or when it contains fluorides), hot concentrated alkali solutions, and superheated water, are capable of etching glass. HF is the most powerful of this group, etching any type of silicate glass. Other agents have significantly lower etch rates and require aggressive conditions and hence are generally not considered for controlled

52


etching. The etching due to HF is attributed to the presence of fluorine containing species F-, HF, HF2- and is depicted by the following chemical reaction: SiO2 + 6HF H2SiF6 + 2H2O

(2.1)

Of these radicals, the bifluoride (HF2-) is considered to have the greatest effect on the etch rate [39]. Etch rate has been shown to increase with HF concentration in the etchant mixture and vary with the inclusion of alkali oxides in its composition [40]. In contrast to glass, quartz has crystalline structure and hence exhibits anisotropic etching much like silicon with the same etching mechanism as described above for glass [41]. Z-cut (Z axis is the optical axis in quartz crystal and the unit lattice is symmetrical around this axis) wafers are used as the etch rate in that direction is greater that any other direction. The etched profiles vary greatly with process conditions, most notably the etchant concentration and temperature. Much like silicon, quartz has several crystalline planes making prediction of anisotropic etch profile difficult. However, several experimental [42] and theoretical [43, 44] studies on etch rates and profile development have been performed and can be used to design masks for obtaining appropriate shapes required for microfluidic structures. HF wet etching in conjunction with various other microfabrication techniques such as metallization, chemical mechanical polishing, and bonding has been used widely to fabricate microfluidic devices. For example, Lagally et al. [38] used wet etching of glass substrates to fabricate an on-chip PCR chamber. As shown in Figure 2.9 the glass substrate was coated with a thin layer of poly-silicon, which served as an etch mask, using a chemical vapor deposition process. The poly-silicon layer was patterned using a CF4 plasma dry etching with a photoresist layer patterned using photolithography as the etch mask. Another popular masking material is metallic multilayers such as Chromium/Gold (Cr/Au) that is sputtered or evaporated on glass substrates and patterned using photolithography [33]. Gold, due to its inert nature, serves as a suitable mask in preventing HF attack on the glass underneath. Chromium serves as an adhesion layer in preventing delamination of Au from the surface. In some instances the photoresist that is used to pattern the Cr/Au layer is left on during etching as additional protective layer to prevent etching through pin holes in the metal layer. Alternatively, thick negative photoresists such as those based on novolak resin or SU-8 have also been used as etch masks through a single photo-


53

lithography step, thus reducing the complexity and cost associated with fabrication [45].

Fig. 2.9 Process flow for fabrication of microchannels in glass.

This patterned substrate was subsequently etched in a 1:1:2 HF:HCl:H2O mixture at an etch rate of 6 µm/min to obtain a final etch depth of 42 µm and a channel width of 100 µm. Soda lime glass is a multi-component mixture consisting of small amounts of Na2O, CaO, MgO and Al2O3 apart from SiO2. Etching of this material using concentrated HF or buffered HF solutions causes precipitate formation which mask subsequent etching of the underlying regions leading to increased surface roughness. Addition of HCl was found to dissolve these precipitate and produce a smoother finish [46]. Subsequently, the photoresist layer was removed and the remaining poly-silicon etched using a CF4 plasma dry etch. Access holes (with diameters of 2.5 mm and .75 mm) to the microchannels were drilled using Diamond tipped drill bits to a depth of 1 mm. The etched and drilled substrate was thermally bonded, at 560 °C for 3 h, to a 210 µm-thick flat glass wafer of identical radius in a programmable vacuum furnace. Tygon tubing of 1/8 inch outer diameter was used to for fluidic connections.

54


2.3.1.2 Plasma dry etching of glass

The working principle of RIE and DRIE for glass is similar to that used for Si. However, the process is complicated due to the multi-component nature of glass. Various gas phase etchants such as CHF, CHF3 and CF4 that produce fluorine free radicals in the plasma and are directionally accelerated to the substrate are used for RIE etching [47]. The resulting fluorine radicals react with silica and produce volatile reaction products in the form as shown in the equation: SiO2 + CF4 CO2 + SiF4

(2.2)

Etch rates for soda lime glass of about 60 Å/min at 300 W and 60 mTorr pressure of CF4/Ar were obtained [47]. The etch rate was shown to vary with the chemical composition of the etchant mixture and the nature of the glass. This relatively slower rate of etching is said to be due to deposition of non-volatile compounds formed during reaction with glass components [48]. Higher etch rates of 0.2 - 1 µm/min for quartz and 500 nm/min for Pyrex have been obtained using inductively coupled plasma with SF6 as the gaseous etchant at pressures of 2-10 mTorr and a high bias of 340 V [49, 48]. 2.3.1.3 Photo-definition of glass

Certain types of glasses can be photostructured using UV light such that they crystallize in the exposed regions and the etch rate is increased significantly. In this way anisotropy induced by UV [50] or laser exposure can be used to fabricate high aspect ratio structures. Such glasses, also known as photoetchable glass, typically contain traces of rare earth metal oxides in them [50]. An example is a lithium aluminum silicate glass, also known commercially as FOTURAN (Schott Glassworks) that contains CeO2, Ag2O and Sb2O3 as rare earth impurities. When the glass is melted to form the wafers some Ce3+ ions are formed that are stabilized by Sb2O3, or other reducing agents: 2 Ce4+ + Sb3+ ↔ 2 Ce3+ + Sb5+

(2.3)

2

On UV exposure at 310 nm (dose of 2 J/cm for 1 mm thick plate) the Ce3+ ions oxidizes reducing to Ce4+ and releasing an electron: Ce3+ + hν (312 nm) Ce4+ + e-

(2.4)

This electron reduces Ag ions to Ag metal atoms: +

Ag+ + e Ag

(2.5)


55

Subsequent heat treatment at 500 °C causes agglomeration of silver atoms and formation of larger nuclei. The temperature is then increased to 600 °C when the surrounding glass region crystallizes around the nuclei forming lithium-metasilicate Li2SiO3 having a different chemical and physical properties than the unexposed glass region. The photodefined substrate is etched using 10% HF solution. An etch rate of ~10 µm/min; a selectivity of ~20; and a minimum line width of 25 µm with aspect ratio of 20 were demonstrated with a roughness less than 1 µm [50]. 2.3.1.4 Fluidic interconnect fabrication

A number of methods are used for creation of fluidic feedthroughs to connect external macrodevices such as pumps and valves with the microfluidic network. They include ultrasonic machining, laser micromachining, abrasive powder machining, micro electrochemical discharge machining, and conventional drilling [51]. The simplest method of machining glass is using single crystal diamond or poly crystalline diamond tools in ductile mode machining at low depth of cuts to precisely define microchannel shapes or for through hole machining. Brittle materials such as glass are difficult to machine mechanically due to fracture generation, tool breakage, and wear. However, under certain controlled conditions such as low depth of cut, it is possible to machine brittle materials such as ceramics using single or multipoint diamond tools so that material is removed by plastic flow, leaving a crack-free surface [52]. The transition from a brittle to a ductile mode can be thought of as a balance between the strain energy and the surface energy to cut a chip. In brittle materials the density of defects is small and therefore when the thickness of the chip cut is small, the stress field generated is also small that it does not encounter any defect and thus does not generate fracture. Poly crystalline diamond tools with randomly distributed sharp protrusions of diamond with dimensions around 1 µm have been used to machine grooves, pockets, and through holes in soda lime glass. The tools are made of a shank of tungsten carbide which is machined using a wire EDM method with surface roughness of 0.3 nm. Abrasive powder machining for glass in the microscale was initially developed at Philips research laboratories for through hole interconnects in glass sheets for flat panel displays [53]. The mechanism is based on material removal due to generation of micro-cracks by sharp indenting particles [54]. The process involves three basic steps. The first is masking using electroplated metals [55] and abrasion resistant polymers such as urethanes

56


[56] to protect the regions that are not targeted for etching. The second is erosion of the exposed parts of the wafer by high velocity jet of abrasive particles. In this step, the size of the abrasive particle has to be chosen small enough to enter the mask openings for effective erosion of the substrate. The size of the abrasive particles, along with the other parameters such as kinetic energy of the particles, determines the depth of cut [57]. The abrasive particle dispensing system provides a constant flow rate of particles with constant kinetic energy uniformly over large surface area. The third step is removal of the masking layer and cleaning of the substrate. Microfluidic channels of 100µm width have also been machined in glass [58] with 30µm alumina particles and resolutions down to 10µm have been achieved with smaller abrasive particles [59]. Electrochemical discharge machining (ECDM), otherwise also described as electrochemical arc machining, spark-assisted chemical engraving, and electrochemical spark machining, is a low-cost method to obtain deep anisotropic features with surface roughness similar to that obtained by HF etching. In this process, the substrate to be machined along with the two electrodes (cathode and anode) are immersed in an electrolyte solution (typically, sodium hydroxide, or potassium hydroxide). The cathode which is connected to the tool is placed close to the substrate. The anode, typically a platinum wire, is placed far away and has a larger surface area than the tool electrode. When the applied voltage is higher than the critical voltage [60], electrochemical reaction results in generation of H2 bubbles at the cathode, temporarily creating a gas film, separating the cathode from the electrolyte. This results in the buildup of voltage across this film and electric discharge across it. The etching is due to the thermal effects of the discharge combined with the chemical corrosion due to the electrolyte accelerated at that spot. Use of a rectangular pulse and tool rotation was found to improve the machining quality and obtain straight sidewalls. Rectangular pulse voltages of 40 V with tool travel rate of 1000µm/min and rotation of 1500rpm has been used to machine grooves of 367µm depth and 200µm width in pyrex glass [61].

2.4 Wafer Bonding For the construction of complex devices it is sometimes necessary to assemble individual layers for the final device. Planarization and whole wafer bonding methods can be used to assemble individual layers to obtain three-dimensional structures that are thicker than a single wafer and this


57

can be done simultaneously for several devices. In microfluidic devices, bonding is used primarily to provide a top cover to open microfluidic channels and networks formed using either silicon or glass using bulk micromachined methods. The other substrate to be bonded could be of silicon, glass, or even other metals and can be instrumented with electrodes, heaters, sensors, detectors etc. to increase the functionality of the device. Several methods have been used for this purpose, including fusion bonding, anodic bonding, and adhesive bonding methods. Bond strength, maximum pressure to failure, low chemical reactivity of interface materials, and high temperature stability are the key factors in the choice of approach. A good review of wafer bonding techniques and metrology is described elsewhere [62] 2.4.1 Fusion bonding Several processes have been developed, with fusion bonding being the most common [16]. In wafer fusion bonding, the silicon wafers are bonded at room temperature where they adhere to each other by hydrogen bridge bonds that develop between the OH terminated surfaces across the interface. The energy of this bond is low compared to that of covalent bonding. The wafers are then annealed at temperatures between 700°C to 1100°C that causes the Si-O-Si bonds to react [16]. Initially as the temperature is ramped up, water molecules at the interface rearrange and produce stable hydrogen bonding at temperatures below 110°C. Further increase in temperature forms stable siloxane bond at the interface at temperatures between 150 and 800°C. Finally, viscous flow of interface oxide occurs at temperatures higher than 800°C. Tensile strengths as large as 9MPa have been obtained using this method. A major problem with wafer fusion bonding is the presence of non-contacting areas on the silicon wafers, which can be caused by particles, residues, and surface defects. Therefore, fusion bonding requires that surfaces are perfectly smooth as small particles can cause a large void [16]. Another major issue is incompatibility of electronic circuits and semiconductor detector that have been fabricated on the wafers to high temperatures that occur in this process. 2.4.2 Anodic bonding Anodic bonding is one of the most widely used methods for bonding of glass microfabricated substrates to silicon, metal, or other glass wafers. The term was coined by Wallis and Pommerantz in 1969, when they observed that by applying an electric field a bond between metal foil and

58


glass could be achieved at relatively low temperature [63] compared to fusion bonding. This is ideal for bonding substrates containing electronic circuits, sensors and actuators or substrates with large difference in thermal coefficient of expansion. Typical operating conditions are 200-1000V and moderate temperatures [300-450°C]. This method has extensively been used in microfabrication of microfluidic devices such as micropumps and microsyringes [64, 65], as well as in other MEMS applications like inertial sensors [66] and pressure sensors [67]. In microfluidics, the microchannel network is sometimes structured in glass and bonded to a silicon wafer using anodic bonding. At elevated temperatures of 300-4500C, the mobility of ions in the insulator is increased [68] resulting in transport of mobile Na+ ions away from the interface. This causes formation of a depletion zone with excess O- ions that are chemically bonded at this elevated temperature to the Si ions in the adjoining wafer [69, 70]. The electrostatic attraction between the two wafers at high electric fields pulls together the interface [71] and hence the process is more tolerant of small surface roughness compared to fusion bonding. The same mechanism explains bonding at the glass-glass and glass-metal interfaces [72]. Glass-glass bonding involves additional step of creating an intermediate layer using dissolved wafer process where a silicon wafer is bonded first to a glass wafer and then etched back to retain a SiO2 layer. This modified wafer is bonded to the structured glass substrate using the same process as described for glass-silicon bonding. Alternatively a layer of silicon nitride, oxide, or carbide can also be used instead of the dissolved wafer process for creation of the intermediate layer. 2.4.3 Adhesive bonding This technique of using an intermediate adhesive layer to bond wafers of similar or dissimilar materials has been in existence for a long time. Typical intermediate layers are polymers such as polyimides, epoxies, thermoplast adhesives, elastomers such as PDMS and photoresists. They are spin coated to one of the substrate to produce a thin uniform layer prior to bonding. Then depending upon the adhesive material relatively low curing temperature (100°C) and moderate pressure is applied to bond them [73]. The main advantages of this technique over anodic and fusion bonding are: low cost, low bonding temperature and greater tolerance to surface roughness and topography. This process does not produce hermetically sealed structures, the life span of polymers is low, and polymers are not suitable for high temperature processes [74]. Formation of voids due to entrapped


59

air and solvent evaporation can also be an issue [75]. The main factor in establishing good bonding strength is absence or reduced release of reaction products in the vapor form during the curing process. In this context PDMS prepolymer has been used as the intermediate layer and transferred on to the bonding surface using imprint lithography [76].

2.5 Polymers Polymers are macromolecules with a high molecular mass and a molecular structure consisting of repeated monomeric units. They can be classified into two categories, linear and cross linked polymers, depending on the position of the reactive groups in the monomer and the cross-linker. The polymerization process is statistically dependent and hence a range of polymer chain lengths with cross-linking is created forming a large 3-D network. Hence polymers usually do not have a defined melting temperature but soften over a range called the melt interval. A wide variety of polymers have been created from a number of starting monomers having different physical and chemical properties [77]. Polymers, depending on their response to thermal treatment, have been classified as thermoplastic, thermosetting, and elastomeric polymers. Thermoplastics are mostly linear or branched polymers that are melted upon the application of heat and re-solidify when cooled. They do not have cross linking and hence their thermal behavior is reversible. At a temperature above the glass transition temperature these materials become plastic and can be molded into specific shapes, which they retain at lower temperatures. Thermosetting polymers are heavily cross linked and hence the molecular movement for elasticity is not possible. They are rigid and brittle and do not soften significantly with increased temperature. Elastomers are weakly cross linked polymers. They can be easily stretched and will revert back to their original shape upon release. Since they are cross linked they do not melt before reaching their decomposition temperature. 2.5.1 Microfabrication Although silicon and glass have been the materials that have been primarily used for microfluidic devices, they have a number of disadvantages. Silicon based devices have high substrate electrical and thermal conductivities and therefore modification in design and additional processing steps are typically required for their use in microfluidic systems. (For ex-

60


ample, insulation layers are needed to prevent a short circuit through silicon substrate in electrokinetic devices. Similarly, isolation islands have to be incorporated in the design for microreaction chambers to prevent thermal cross talk. Glass provides excellent isolation properties but is very difficult to machine into high aspect ratio complex 3-D structures. Polymers in contrast are moldable to complex shapes and dimensions and have comparable isolation properties to glass. Polymers also have a number of attractive properties such as low cost, chemical inertness, low electrical and thermal conductivity, suitability for surface modification and compatibility with biological materials that make them ideally suited for use in biological microfluidic applications [78]. Some of the common thermoplastic polymers include polymethylmethacylate, poly-carbonate, poly-vinylidenefluoride, poly-sulfone, poly-styrene, poly-vinylchloride, poly-propylene, poly-etheretherketone, polyoxymethylene, and poly-amide. These polymers cannot be microstructured using the traditional microfabrication techniques. However other techniques such as injection molding, hot embossing, and casting can be used for microfabrication of polymers by exploiting their compliant form at high temperatures. In these methods, a substrate made of the polymer is fashioned as a negative replica of the master, which retains its structural integrity at these temperatures. Once a master mold has been made, typically using traditional microfabrication techniques, several thousand polymer replicas can be molded with low cost. The replication process is insensitive to the complexity and fineness of the design, and can be performed outside the clean room while still obtaining device resolution similar to photolithography. This aspect of polymer micromachining reduces the cost of the devices significantly – making the technology ideal for disposable biomedical devices [79]. Surface micromachining techniques have also been recently adapted to suit low temperature processing for polymeric materials. These allow construction of devices with multiple materials and complex functionality. Outlined below is a brief description of the different methods widely used to structure 3-D objects in the microscale with polymer substrates. Several companies such as Micronics, Steag microParts, MicroTec, Amic AB, Micralyne, and DALSA provide foundry services for fabrication of microfluidic chips in polymers. 2.5.1.1 Injection molding

Injection molding has been one of the first techniques adopted for micro manufacturing of polymeric structures for microfluidic devices. It is a well established technique and the manufacturing infrastructure, technology and


61

process know-how already established have been rapidly adapted for microscale molding [80]. Several groups have demonstrated the feasibility of replication in thermoplastic polymers of microstructures [81, 82, 83, 84]. Similarly, a number of investigations have also been performed on fabrication of micromold inserts with microscale features. These include using microfabrication techniques such as silicon micromachining [9], SU-8 photolithography, [85] and other machining techniques such as micro EDM and LIGA process [86]. In injection molding the mold cavity with the mold inserts is evacuated and heated to a temperature higher than the glass transition temperature of the polymer. A horizontal injection unit heats the polymer and injects the viscous melt into the mold cavity. The entire cavity is then cooled below the glass transition temperature and subsequently the molding part is demolded. Figure 2.10a depicts the operation. The quality and precision of the mold insert determines the quality of surface features. Injection molding is the most suited for industrial production due to short cycle times (~ mins). 2.5.1.2 Hot Embossing

Hot embossing refers to a stamping process where microstructural features from a hot mold insert (master) is transferred on to a thermoplastic substrate. Figure 2.10b depicts the process. In this method, a thermoplastic film or substrate is inserted into the embossing mold with mold inserts on either side. The mold is heated to a temperature just higher than the glass transition temperature of the thermoplastic material. The tool is evacuated and the heated mold inserts are pressed against the thermoplastic film with high force. The thermoplastic material close to the surface, due to the mechanical force and temperature is redistributed in such a way as to form a negative replica of the feature on the mold insert. The setup is cooled and the mold insert is withdrawn from the plastic [79]. In injection molding, the material is injected and fills the cavity at high temperature. Subsequent cooling introduces significant stresses in the bulk of the material. Unlike injection molding, in hot embossing the thermoplastic material flows only over short distances, which results in low stress. Local heating and a low thermal capacity requirement also reduces the thermal time constant for embossing and delaminating and hence achieves a faster process time [79]. Since high aspect ratio structures (>10) can be replicated quite easily in thin films (1-100 µm), this technique has been adapted to replicate patterns on photosensitive films using a process called nano-imprint lithography.

62


Fig. 2.10 Process flow for various fabrication processes available for microstructuring polymers.


63

2.5.1.3 Casting

In this process a material is introduced into a mold in its liquid state, allowed to solidify in the shape inside the mold, and then removed producing a replica of the microstructured master. Figure 2.10c illustrates the process. Molds for casting are also created similar to those for injection molding and hot embossing. A number of materials including hot, liquid metals and thermoplastics have been cast. A variation of this process is what is termed as reaction casting. In this process a two component mixture consisting of a long chain polymer (base) and short chain polymer with initator (crosslinker) is used. The mixture in the uncrosslinked state is fluid and wet the surface attains the shape of mold insert when cast. Subsequent exposure to heat or UV light input coldsets the mixture through cross linking - increasing its viscosity by several orders of magnitude. This process is typically used for non-thermoplastic materials such as thermosetting plastics (bakelite), epoxies [87], and elastomers (PDMS) [88]. In this process the drawbacks are that complete mixing and distribution of the two components have to take place and the chemical reaction (cross linking) has to take place throughout the bulk for uniform strength. This can only achieved by comparatively long cycle times (~hours). However feature in the scale of a few nm can be easily replicated. 2.5.1.4 Stereolithography

Stereolithography is a rapid prototyping technology first developed in 1980’s [89] for prototyping in CAD/CAM applications. Figure 2.10d shows the schematic of the process. Here, a laser beam is focused on the free surface of a photosensitive liquid to induce polymerization of the local region of liquid and to transform it to a polymerized solid. The object to be formed is deconstructed into series of two-dimensional layers and reproduced by laser induced polymerization of a resin. Resolution depends on the laser wavelength, scanning apparatus used and the diffusion in the photosensitive liquid. Penetration depth of the laser light into the liquid determines the thickness of the layer. After laser exposure of one 2D crosssectional layer shape, fresh liquid is spread on top and the second layer is created similarly. The resolution of this technique is ~150-200 µm in three dimensions. This resolution has been improved with the use of a glass holder (through which the laser light transmits) attached to an X-Y-Z micro-positioner to accurately control the position and thickness of the resin with which the light interacts [90, 91] in the resin volume. Another method, free surface polymerization, was then developed wherein the part formed is attached to the X-Y-Z positioner and moved close to the free

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surface of the resin/ air interface [92] achieveing micrometer resolutions. The throughput has been increased by using optical fibers to split the laser light and to manufacture several part simultaneously [93]. Liquid Crystal Display (LCD) devices [94] and Digital Micromirror Devices (DMD) [95] have been used to generate patterns and eliminate X-Y movement and provide a lithography type process which decreases the time to generate layers and increases throughput. An excellent review of stereolithography and its applications has been published [96]. 2.5.2 Polymer materials: Some of the most popular and widely used polymeric materials in microfluidics include Parylene (paraxylylene), PDMS (polydimethylsiloxane), SU-8, hydro-gels, acrylate based porous polymer monoliths, biodegradable polymers, polyesters, polyimide, and Paraffin. Some of these polymers such as parylene, PDMS, SU-8, biodegradable polymers, and polyimide are primarily used for the construction of microfluidic channels. Others including paraffin, hydrogels, and porous monoliths are used as functional material in the construction of valves, pumps and reactor beds. This section deals with the detailed description of the properties of these polymers and their fabrication processes. 2.5.2.1 Parylene

Parylene is the industry name for a class of polymers called poly paraxylene. It is available commercially in three dimeric stable forms, which are variations of a basic polymer backbone of xylylene with replacement of 1-4 atoms in the benzene ring with chlorine as shown in Figure 2.11. It has been used for a wide variety of applications such as encapsulation for microelectronic circuits [97, 98], as interlayer dielectrics [99], and for microchip packaging [100]. Similarly, a number of applications of parylene in microfluidics include microchannels [101, 102], microvalves [103, 104, 105], membrane filters [106], and other micromachined devices. Parylenes have to be polymerized in-situ and cannot be formed by extrusion or molding techniques because of their high molecular weight. Parylene polymers are currently deposited by vapor deposition and in-situ polymerized at room temperature and in vacuum. The dimer form of parylene, which is solid at room temperature, is sublimated at 140o-160oC and passes as a vapor into the pyrolysis chamber where it then splits into two monomers at 680oC. The reactive monomer flows into a deposition


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chamber where it deposits and begins to polymerize on surfaces below 100oC. The deposition pressure is ~ 100 mTorr and hence the deposition is conformal. This allows for the formation of uniform conformal thin films.

Fig. 2.11 Molecular structure of various commercially available parylene polymers.

Parylene thin films are extremely conformal even with high aspect ratio structures due to the vapor based deposition process. Low surface roughness, pinhole free coating, and excellent dielectric breakdown characteristics can be obtained for thickness greater than 0.5 µm. The films also are chemically pure due to lack of initiators and catalysts in the polymerization process. They are stress free due to room temperature polymerization process. The polymerized films are clear and possess low auto fluorescence allowing visualization of biological and chemical interactions through optical means. They are chemically and biologically inert and biocompatible. Hence they are resistant to damage by acids, bases, corrosive

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body fluids, electrolytes, enzymes, and proteins. Parylenes have superior barrier properties compared to silicones and other polymeric materials. Their moisture vapor permeability is 1.7 x 10-16 kg-m/N.s., which is an order of magnitude lower than silicones. They resist room temperature chemical attack and are insoluble in all organic solvents up to 150° C. Parylene C can be dissolved in chloro-napthalene at 175° C, and parylene N is soluble at the solvent's boiling point (265° C). Substituted parylenes have been developed recently that can react with biomolecules to promote strong attachment [107]. A surface micromachining process for construction of Parylene microchannels has been demonstrated [108]. The process begins with the initial coating of the substrate with a thin conformal parylene layer. Adhesion to the substrate is usually obtained with plasma treatment and silanization by vapor deposition. Next, electrodes and other functional parts of the microfluidic device can be deposited and patterned. Thick (20 µm) photoresist is spun cast and photo-lithographically patterned to define the microchannel shape and dimensions. A second layer of parylene is then conformally deposited to form the top and sides of the channel. The adhesion of this layer is also assisted by a short oxygen plasma treatment. Using a thick photoresist mask, reservoir, and contact pad openings are etched using oxygen plasma. Dry etching in O2 and CF4 atmospheres at low pressures produces etch rates of 0.2 µm/min at 250 W and 50 mTorr [109]. The sacrificial photoresist in the microchannel is then dissolved in acetone for 36 hours without agitation and rinsed and dried. The release time could be shortened by constant agitation or heating the acetone. A number of devices such as DNA separation devices [108], PCR systems [102], cochlear implants [110], microvalves for drug delivery [104, 105], and microneedles have been fabricated using these techniques 2.5.2.2 SU-8

SU-8 was developed by IBM [111] for LIGA applications. A modified version of this material was used to produce high aspect ratio structures for MEMS applications using conventional lithography [112]. Although positive photoresists have been used widely as sacrificial layer for surface micromachined construction of microchannels [102], SU-8 was one of the first negative tone resists to be used as a structural layer for MEMS and microfluidic applications. It consists of EPON SU-8 resin (an epoxy) as its main constituent with γ-butyrolacton as the solvent and 10% wt of photoinitiator. The versatility of this resist was demonstrated by the fact


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that thicknesses from 750 nm to 450 µm can be obtained [112]. It is commercially available from MicroChem Inc. Processing and lithography of SU-8 is similar to conventional negative tone photoresists. SU-8 is spun on silicon wafer and pre-baked at 95oC for a duration of 15 min for every 100 µm thickness. The resist layer is exposed in a UV aligner at a dose of 300-400 mJ /cm2 at 365 nm per 100 µm thick resist layer. The resin has very low UV absorption and hence uniform dose and crosslinking throughout the bulk of the resist can be achieved. It is developed using propyleneglycol monomethylether acetate solution and rinsed in iso-propyl alcohol. The resist can be stripped at this stage using hot 1-methyl-2-pyrrolidon solution. Further hard baking of the resist at higher temperatures 120-200oC for 30 min makes it resistant to metal etchants and other solvents typically used in microfabrication [112]. High aspect ratio structures (>18) with a range of thickness from 80-1200 µm, with conventional mask aligners have been demonstrated [113]. Higher aspect ratios (>60) have been achieved using x-ray sources for LIGA applications [114]. Another feature of SU-8 is that taller structures can be constructed by using multiple layers. A 100 µm layer can be spun, pre-baked and exposed and this can then be used as a substrate for the next layer. Multiple layers with alignment using different masks can produce complex 3-D structures. After the final layer has been exposed and baked, all the layers can be developed at the same time. Several methods for fabrication of microchannels using SU-8 have been reported as depicted in Figure 2.12. A simple method as depicted in Figure 2.12a is the use of a sacrificial positive tone resist to define the microchannel structure. SU-8 is cast over this sacrificial mold and the access holes are lithographically defined. Then the sacrificial resist is removed to create an open microchannel [115]. A variation of this method is depicted in Figure 2.12b. Here, a layer of SU-8 is spun, pre-baked and exposed into patterns of microchannels. Then a thin metal layer is deposited on top to protect unexposed regions from exposure in the bottom SU-8 layer. A second SU-8 layer is then spun, pre-baked and exposed to a pattern to create access holes for the microchannels. Subsequently the second SU-8 layer is developed, metal layer etched and the unexposed resist in the first SU-8 layer also developed to produce the channel structure. [115]. A third method as shown in Figure 2.12c, depends on the energy of proton beam to define the penetration and hence exposure depth of the resist to produce three dimensional features. Initially a thick layer of SU-8 is spun cast. A

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long exposure to 2MeV proton beam through a mask is done to produce a pattern of the microchannels throughout the entire depth of the resist layer that defines the walls of the microchannel. Subsequently the wafer is flood exposed to 0.6 MeV beamwith a penetration depth of 10 µm that polymerizes the entire top layer forming the top surface of the microchannel. Subsequently the unexposed regions are developed to produce the embedded microchannel structure [116]. The fourth method is depicted in Figure 2.12d. Here, the SU-8 layer is spun, pre-baked, exposed, and developed to produce a pattern of the microchannel. A dry Riston photoresist film is laminated over the SU-8 layer. Access holes are then patterned in the Riston layer to provide fluidic connection to the SU-8 microchannel [117].

Fig. 2.12 Methods for fabrication of microfluidic channels in SU-8.


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2.5.2.3 PolydiMethylsiloxane (PDMS)

PDMS is one of the most widely used polymers in microfluidics and BioMEMS. It belongs to a class of polymers called silicones containing a Si-O backbone. To this backbone, organic groups are frequently attached to the silicon atoms via a Si-C bond. PDMS has a repeating (CH3)2SiO unit. Depending upon the number of repeat units in the polymer chain and the degree of cross-linking, the material can be obtained in different rheological forms such as fluids, emulsions, lubricants, resins, elastomers, and rubbers [118]. The low rigidity of the backbone allows the methyl groups to be easily exposed, resulting in low intermolecular interactions of PDMS and also its low surface tension. PDMS in its elastomeric form is used generally for microfluidic and BioMEMS applications. The commercially available version is provided as a kit (Sylgard 184 from Dow Corning and RTV615 from GE) with the PDMS base solution and a curing agent to cross link siloxane base oligomers (in the base) containing vinyl terminated end groups (CH2=CH-) into elastomer by an organometallic crosslinking reaction. The base solution also contains a platinum based catalyst and silica filler. Typical crosslinkers are dimethyl methylhydrogen siloxane or tetrakis(dimethylsiloxyl silane) (TDS). The curing agent also contains an inhibitor (tetramethyl tetravinyl cyclotetrasiloxane). The platinum-based catalyst catalyzes the addition of the SiH bond across the vinyl groups, forming Si-CH2-CH2-Si linkages. The multiple reaction sites on both the monomer chain and crosslinking chain enable three-dimensional crosslinking. A casting process is used to form microfluidic channels and structures using PDMS elastomer as the structural material [119]. The master molds for this process with high aspect ratios are created using wet chemical etching of lithographically patterned silicon substrate. Alternatively, SU-8, and poly-urethanes have also been used to form the master molds [120]. A mold release layer of 3% (v/v) dimethyloctadecylchlorosilane in toluene is vapor deposited for 2 h to facilitate delamination of the PDMS replica. The PDMS base and its curing agent, mixed in 10:1 ratio, are cast over the mold. The PDMS pre-polymer mixture, due to its low surface energy, wets the entire surface down to the molecular level forming an accurate negative replica of the relief. Curing of the PDMS prepolymer for 4 h at 65 oC initiates crosslinking reactions, which solidify the material into an elastic monolith. Subsequently, the PDMS replica is peeled off the master. The master can be used to create several PDMS molds, thus replicating micro and nanoscale features without the use of expensive clean room processes.

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Access holes to reservoirs ae punched through the bulk material using a sharp needle. The device is then placed on a thin slab of PDMS in order to form a closed channel system of four equivalent walls. A hermetic seal was obtained by mere contact without applying large external force [119]. The chips manufactured using this technique withstood pressures of 1 bar. Buffer solution and separation media were pipetted into the reservoirs, and the microchannels were filled by applying vacuum to one of the reservoirs. Capillary electrophoresis separation of φX-174/HaeIII DNA was accomplished in this device, to demonstrate the feasibility of microchannel construction and its use in BioMEMS [119] Although adhesion between PDMS layers was obtained in the previous process, the bond was reversible. A method to irreversibly bond PDMS to a substrate in order to construct devices to withstand higher pressure has also been developed [120]. In its natural state the PDMS surface is inert due to hydrophobic CH3-groups on the surface, resulting in low adhesion. Surface oxidation of the PDMS in plasma removes some of the methyl groups and exposes the PDMS backbone containing hydroxyl groups. Oxidized PDMS surfaces brought into conformal contact with each other form a tight, irreversible seal, improving adhesion of the substrates and pressure tolerance of microchannels. The Si-OH groups at the surface of the two PDMS slabs form a covalent Si-O-Si bond between them. A prolonged plasma treatment (>30 seconds) however, results in the oxidized layer becoming thinner and cracking. If the treated samples are left to age in ambient atmosphere the hydrophobicity recovers due to low molecular weight PDMS migrating to the surface [121]. Although plasma oxidation is most popular, other techniques such as corona discharge [122] and UV light exposure [123] have also been used to reduce hydrophobicity, particularly to reduce protein adsorption in microchannels. Microvalves and pumps have been incorporated into PDMS microfluidic networks by modifying the fabrication technique described above to produce multiple stacked layers with microscale features embedded in them using a technique known as “multilayer soft lithography” [124]. In this process, several elastomeric layers consisting of microchannel networks and PDMS membranes are fabricated separately, aligned, assembled and bonded. This technique exploits the two-component curing process to obtain bonded PDMS interfaces. The bottom layer has an excess of one of the components (PDMS base), whereas the upper layer has an excess of the other (crosslinker). When these two layers are brought in contact with each other and thermally cured, crosslinking reaction occurs between excess materials in these layers forming covalent linkages and a hermetic


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seal. The strength of the interface equals the strength of the bulk elastomer [124]. Additional layers are added by simply repeating the process with the polarity of material excess reversed. Valves and pumps using the pneumatic control have been demonstrated by constructing a two microfluidic layers sandwiching a membrane layer with the top layer being the pneumatic control network and the bottom layer being the fluidic network [124]. 2.5.2.4 Polyimide

Polyimide has also been used as a structural material for the construction of microfluidic devices such as microvalves and pumps [125, 126] due to its excellent chemical and thermal stability, low water uptake, and good biocompatibility. Commercially available polyimide resin consists of aromatic heterocyclic chains of alternating carbonyl groups, which act as electron acceptors and nitrogen groups which act as electron donors. This charge transfer complex stabilizes the chain and is responsible for its chemical inertness, structural rigidity, and high strength. Polyimides are made photosensitive by attaching a photosensitive group that is an inhibitor of polymerization. The photosensitive group detaches from the backbone upon UV excitation, initiating polymerization in that region [127]. A photosensitive commercially available polyimide (PI – 2732, Dupont) has been used as structural material for microchannel construction [128]. In this process, a thin polyimide layer is spun, pre-baked and patterned on a substrate using optical lithography with the desired geometry and vent channels, but not cured. Next, a thin layer of solvent with dissolved precursor is used to coat a soft-baked layer polyimide, on a second substrate. The two halves are placed in contact and cured. Channels with dimensions 50 to 1000 µm wide, 3 to 30 µm deep have been fabricated for chemical analysis and heat transfer devices [128]. The structures developed in this process have significant internal stress due to solvent evaporation during the final curing. To resolve this problem, a modified lamination method has been developed to create microchannels with embedded electrodes [126]. This lamination process uses a combination of photosensitive polyimide (PI – 2732) that is microstructured using optical lithography and non-photosensitive polyimide (Mylar film), which is microstructured using dry etching with oxygen plasma and a silicon dioxide mask. The PI-2732 polyimide is spun cast to obtain a 5 – 20 µm layer, exposed and cured at 350oC for 1 hour. This layer forms the base of the microchannel. A layer of Ti/Pt (50 nm /

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200 nm) is sputtered and patterned using a positive photoresist. This forms the electrodes inside the microchannels. A second layer of PI – 2732 polyimide (5 – 20 µm), which forms the side walls, is spun, exposed and partially cured at 100-150oC for 1 hour. This layer not only insulates the electrodes but also provides the structure for microchannels. This layer is treated with n-methyl-2-pyrrolidone (NMP) (swelling agent) that causes the partially cured second layer to swell, enabling higher interdiffusion with other polyimide layers during lamination. A thin Mylar foil is spin coated with photosensitive polyimide, partially cured, flipped over, and bonded to the substrate by lamination. This layer forms the top of the microchannel. The Mylar layer is coated with oxide, access holes lithographically defined and dry etched in oxygen plasma [126]. No interface was observed between the open channel pre-structures and the laminated top layer, which indicates good channel sealing as shown in Figure 2.13. The channels thus fabricated were able to withstand pressures as high as 19 bar [126]. The curing results in shrinkage of the polyimide (30–40%), which degrades the sharpness of the sidewalls.

Fig. 2.13 Cross section of microchannels created in polyimide. Reproduced with permission from [126] © 2001 Royal Society of Chemistry.

2.5.2.5 Hydrogels

A gel is a loose network of cross-linked polymers that retains much of its solvent. Gels in which water is the solvent are called hydrogels. Any water-soluble polymer can be prepared as a hydrogel by carrying out the polymerization in the presence of a cross-linking agent in an aqueous solution of the polymer [129]. Hydrogels have been recently introduced in microfluidic devices to provide intelligent transduction of signal in one domain (biological, chemical, mechanical or fluidic) to another. Hydrogels can be engineered to be sensitive to a variety of environmental factors such as pH [130], temperature [131], electric field [132], glucose concentration [133], and antigens [134] and convert them to mechanical action such as expansion or contraction.


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Diffusion of the stimuli into the bulk of the gel is the key factor in determining the response time [135]. Therefore, faster response can be achieved by reduction in size, making hydrogels attractive materials for microscale actuation. Since these gels have high solvent content, their properties depend upon the balance of polymer-polymer vs. polymer-water interactions. Structural change, size variation, and mechanical actuation can be accomplished by slight changes in the polymer-solvent interaction properties initiated by external stimulus of which temperature is the most common [136]. A linear polymer that displays cloud point behavior when heated can be crosslinked to give a temperature-sensitive gel network. For example, poly (Nisopropylacrylamide (polyNIPAM) or related copolymers, when heated undergo a rapid and reversible phase transition from extended hydrated chains to collapsed hydrophobic coils and shrinks by expelling water above a temperature termed as Lower Critical Solution Temperature (LCST) of 32 oC. These gels can be formed by photoinitiation, particularly with the use of UV light [137]. This feature makes them suitable for microfabrication through photolithography. A solution of Nisopropylacrylamide (NIPAM), allyl methacrylate and azobisisobutyronitrile (photo-initiator) in tetrahydrofuran (THF) solution was spun cast to 1 µm thickness and photo-lithographically defined using optical photolithography [138]. The solution can also be cast in a sacrificial mold on the substrate to obtain higher thicknesses. Photolithography and development in acetone produces microstructures with 50 µm resolution. The transduction property of this polymer has been used to fabricate an inline microvalve [138]. A 2.5-fold increase in polymer volume in water is observed when the water temperature changes from 25°C to 10°C as seen in Figure 2.14. Response time was reduced to 7 s primarily due to microfabrication of the hydrogel [138].

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Fig. 2.14 Microfabricated hydrogel actuated using temperature.

pH sensitive hydrogels have also been used to regulate flow in microfluidic channels [139]. The photopolymerizable liquid consists of acrylic acid and 2-hydroxyethyl methacrylate (in a 1:4 molar ratio), ethylene glycol dimethacrylate (1 wt%), and a photoinitiator (3 wt%). In this method, PDMS microchannels were filled with the prepolymer and used to provide spatial resolution. This mixture was irradiated with UV light through a photomask to define the regions in the microchannel for polymerization [139]. Polymerization times varied depending on light intensity, photoinitiator, and monomer mixture, and was less than 20 seconds when Irgacure 651 was used as the photoinitiator. When the polymerization was finished, the channel was flushed with water to remove the unpolymerized liquid. In a different implementation, hydrogels responsive to glucose concentration were fabricated using a casting based approach [140] for use in microvalves. The expansion of the gel in response to glucose concentration regulates the flow in the microchannel situated below. This gel, also displays sensitivity to pH and demonstrates a 100% volumetric expansion to pH variation from 7 to 11 and a 30% volumetric expansion for glucose variation from 0 to 100 mM at physiological pH of 7.4. As noted earlier, the response time of the gel is dependent on its size and valve opening times as low as 7 min have been observed with 30 µm hydrogel layers [140]. 2.5.2.6 Macroporous polymers

Macroporous polymers are monoliths whose internal structure consists of an interconnected array of polymer microglobules separated by pores, and their structural rigidity is secured through extensive crosslinking [141].


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These polymers have a fixed pore size, narrow pore size distribution, and due to extensive cross links are structurally stable with their porous structure persisting even in the dry state. These materials were initially produced in bead forms for applications as chromatographic and ion exchange column packings [142]. Phase separation of inert diluents (porogens) during polymerization was used as the mechanism for precise pore size control and its narrow distribution [143, 144]. They have also been demonstrated to be effective as monolith support for catalysis reactions [145]. The fabrication of these monoliths in molded fashion in various tube sizes and tube materials have been demonstrated [146, 147, 148]. These polymers have been used in high throughput bioreactors with immobilization of various catalytic enzymes on the pores [149], and are now finding applications in drug delivery, membranes, and encapsulation of bioreactors [129]. The surface properties of these pores formed can be changed from hydrophobic to hydrophilic by addition of grafted side chains [150]. The porous nature gives them high surface to volume ratio, which is desired in case of catalytic reaction, surface adsorption, chromatography, drug delivery, and electroosmosis. Microfabrication of these monoliths, has been demonstrated by using conventional lithographic techniques [151]. The main issues encountered were the low viscosity of the monomeric mixture and the effect of oxygen on the polymerization reaction. Low viscosity, restricts the ability of using spin coating to obtain defined uniform thickness of films. Very thin layers (~50nm) and non uniform distribution of polymer thickness were achieved by spinning. Secondly, oxygen is an inhibitor in the polymerization process. Thin layers of the monomeric solution exposed to atmosphere absorb oxygen, which inhibits polymerization. These issues were resolved using a microstructured glass mold which was etched to a depth of 20 µm and using it to enclose the monomeric mixture [151]. The substrate on which the polymer monolith is to be structured is used as the capping layer. Adhesion to the substrate was enhanced by depositing a monolayer of gammamethacryloxytropyl trimethoxy silane to it while it is reduced with glass mold by coating it with parylene [151]. Since the glass mold is optically transparent the monolith could be patterned using direct lithography. Light initiates radical generation of AIBN and polymerization of the monomers. Upon polymerization the polymer phase separates from porogen phase and polymerizes around the trapped porogens forming a porous structure. However, low viscosity causes diffusion of polymers and loss of resolution of the features created. Hence flood polymerization and using dry etching techniques for structuring creates high resolution structures as shown in Figure 2.15. These structures have been demonstrated to increase the

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pumping capacity of micro electroosmotic pumps [151] and have potential applications in microscale bioreactors and filtration units.

Fig. 2.15 Macroporous polymers patterned on silicon wafers using conventional micromachining techniques.

2.5.2.7 Paraffin

Paraffins are saturated hydrocarbon mixtures, consisting of alkanes of varying chain lengths. The general chemical formula for straight chain paraffins is CnH2n+2 and have different melting temperatures based on their chain lengths. They have low thermal and electrical conductivity and low chemical reactivity and therefore are ideally suited for microactuator applications. Paraffins are stable up to 250oC, therefore no boiling occurs even at high temperatures. Further liquid paraffins are non-polar and hence do not mix with the polar liquids such as water which are commonly used in microfluidic applications. They undergo phase change from solid to liquid which is accompanied by volumetric expansion. Differences in volume between phases of 5-40% are observed for various paraffins [152]. Paraffins are also stable through numerous phase change cycles and their properties remain constant over long periods of time. Phase change is one of the preferred methods of actuation because of high displacements and forces obtained. Some of the early studies were focused on developing mini paraffin actuators for endoscopic surgery [153]. These studies led to the development of a 130 mm x 2 mm actuator with a 90 mm x 2 mm paraffin chamber. Volumetric expansion of 14 – 16% were achieved with pressure generation of upto 20 Mpa and stroke length of 9-


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10 mm. Non traditional techniques such as screen printing have also been used to pattern paraffin in reservoirs 1 mm in radius and 0.5 mm in height [154]. Several such reservoirs arranged in series functioned as a peristaltic pump generating a flow rate of 74 nl/min. The main problems with this configuration was reported to be long response times (in seconds) and leakage of paraffin due to insufficient adhesion between the actuation membrane and the paraffin reservoirs. Although this early demonstration showed the promise of paraffin as an actuation material, integration of phase change materials into microsystems has been challenging [155] due to the fact that the material has to be introduced during post fabrication stage. This restricts the size of the actuator to upwards of 1 mm or more. Development of microfabrication techniques for paraffins in their solid form and its compatibility with polymer processing techniques has enabled ease of integration and development of miniature high force actuators in the microscale [105]. Paraffins have high vapor pressures and consequently evaporation is the preferred technique for thin film deposition. Deposition conditions of 5 µTorr chamber pressure and 150oC evaporation temperature provided a deposition rate of 100 nm/min. Adhesion of the deposited film to the substrate was enhanced by silanation and oxygen plasma treatment of the substrate prior to deposition. Surface roughness of 42 nm and less have been measured [152]. Pattering of this paraffin thin film followed an elaborate procedure involving 5 steps due to its temperature sensitivity and dissolution in solvents such as acetone and developer that are used in photolithography. Microactuators consisting of a microheater with a patterned micropatch of paraffin enclosed in a parylene sealed chamber have been developed [156]. Deflection of 2.7 µm at 150 mW was observed. Actuators were assembled into a valve format and a 6bit microflow controller was developed which had flow control from 0.1 – 2 sccm using 5 V, 50 mW power [156]. Another interesting use of Paraffin has been as a modifiable, mobile physical separator between microfluidic channels [157]. In this manifestation, the paraffin is introduced into one arm (stem channel) of a T junction microchannel network as shown in Figure 2.16. The other two arms are the inlet and the outlet. The stem channel is instrumented with several microheaters at various locations. The melted paraffin is introduced into the stem channel and flows by capillary action. The paraffin solidifies as it flows in the channel losing heat to the surroundings. Electronic control of the heaters in the stem channel allows precise positioning of the paraffin interface. The paraffin is initially allowed to flow and solidify at the T junction and physically separate the inlet and outlet arm. The valve is

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latched in this position and blocks pressures as high as 250 psi without additional energy input. Subsequently the valve can be opened by activating the heaters, melting the paraffin and applying a pulse of vaccum at the stem channel interconnect. The duration of the pulse and the temperature of the paraffin determine the movement of the paraffin plug.

Fig. 2.16 Schematic of the phase change valve operation. (i) Loading wax by actuating inlet port heater. (ii) Closing valve by actuating the inlet port and stem channel heaters with pressure at inlet port. (iii) Opening valve by actuating the stem channel and intersection heaters with vacuum at the inlet port. (Reproduced with permission from [157] © (2004) American Chemical Society)

2.5.2.8 Biodegradable materials

Biodegradable materials have traditionally been used as scaffolds for tissue engineering applications [158], controlled release mechanism for drug delivery [159], and in sutures [160]. Commonly used biodegradable polymers are polyglycolic acid (PGA), polylactic acid (PLDA), polycaprolactone (PCL) and poly (lactide co glycolide) (PLGA). The degradation times of these polymers range from 5 months to greater than 24 months [161]. These polymers have low toxicity and minimal immune response, tunable degradation rates, and excellent mechanical properties suited for tissue engineering applications. The degradation of these polymers is by hydrolysis of unstable linkage groups in the backbone yielding degradation products that are capable of re-absorption by the body through metabolic pathways. For example, the degradation product of PGA is glycolic acid, which is a natural metabolite. Suitable unstable groups include esters, anhydrides, orthoesters, and amides. The structures of these polymers allow tailoring of their degradation rates. Synthesis of these polymers is carried out from their stable dimeric version. Figure 2.17 shows the chemical structure and synthesis of most common of the biodegradable polymers. The dimer is cyclic and hence


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stabilized. In the presence of heat and an appropriate catalyst, it undergoes ring opening polymerization and forms linear chains. They take the shape of the mold in which the reaction occurs. Other techniques like injection molding, extrusion have been used for shaping these polymers. An excellent review of the material properties of various classes of biodegradable materials has been recently published [162].

Fig. 2.17 Structure and synthesis of common biodegradable materials.

A wide variety of microfabrication methods including imprinting or hot embossing [161], soft lithography [163], direct writing [163], stereolithography [164], and laser micromachining [165] have also been used for structuring biodegradable materials. Photocurable biodegradable versions of PCL have been developed for setereolithographic applications [166].

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Microfabrication of these photopolymers using lithography has also been accomplished [167]. Compression molding combined with low temperature bonding of layers has been used to demonstrate the feasibility of creating microstructures, specifically microchannels, using biodegradable materials [168]. The application of these devices is in tissue engineering of microfluidic vascular networks, which then form the scaffold for seeding and growth of cells to form tissues with pre-existing blood supply networks. Biodegradable materials are attractive since they provide the initial structural framework for growth and then wither away when the tissue is able to sustain itself. The masters for molding containing the fluidic network are created out of PDMS using soft lithography techniques. PDMS posts are then attached to the reservoirs in the mold. Another PDMS master with larger holes that are concentric with the attached post is assembled on the first master. PLGA pellets are placed between these two layers and heated to 110oC along with application of compressive force of 100-500 lbs. The PLGA becomes a viscous melt and conforms to the microstructure of inside of the mold cavity. Film thickness, 100-500 µm, can be obtained by varying the compression time, temperature and applied force. An unpatterned PLGA film is used to close the microchannel structure using low temperature fusion bonding at 60oC for 30-60 min. Bonding at temperature slightly higher than glass transition temperature maintains structural integrity while still allowing interdiffusion in the areas of contact providing higher bond strength [168]. Multichannel stacks could be created this way leading to 3-D microchannel network as shown in Figure 2.18. A similar method was used to mold PCL material into microchambers for drug delivery [169].


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Fig. 2.18 Microfluidic networks created in biodegradable materials. A) Complex branching of network of microchannels (Scale bar 500 µm); B) High Resolution lines (Scale bar 2 µm); C) Nominal microfluidic channels (Scale bar 50 µm); D) High resolution microchannels (Scale bar 2 µm); Reproduced with permission from [168] © 2004 Wiley.

Another interesting technique for creation of a microfluidic channel network for tissues has been the use of cell seeded alginate as the structural material that is casted on a microfabricated mold [170]. In this study calcium alginate is used as the tissue scaffold due to several favorable properties such as its appropriateness for long-term culture and formation of functional tissue, compatibility with macroscale molding to form cellseeded structures with physiological geometries and its ease of chemical modification to enable the presentation of biospecific extracellular ligands [170]. In this process PDMS masters with the appropriate microchannel network features were created using SU-8 lithographic techniques described above. Aluminum molding jigs were created and the master was placed in it. Subsequently cell-seeded alginate (4% (w/v) calcium alginate) was injected and cross-linked. Multiple layers of the molded alginate were sealed together by application of pressure by the aluminum jig. Replication of 100 µm channels have been achieved using this technique [170] with high cellular viabilities. The degree of viability after fabrication was sensitive to the magnitude of shear imposed during the dispersal and injection of the cells in the alginate solution.

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2.6 Conclusion There has been growing interest and effort over the past 15 years towards development of miniaturized and automated devices for chemical and biochemical processes and analysis in areas as diverse as medical diagnostics, drug discovery, specialty chemical production, cell handling, and analytical laboratory applications. A significant majority of microfluidic devices are currently fabricated in silicon and glass primarily due to established fabrication techniques that have historically been developed for these materials. A further advantage is that most of the biochemical reactions have been characterized and standardized in glass. The use of polymers in microfluidics has been recent but is rapidly growing. A number of microfabrication methods, both conventional and non-conventional, have been developed to encompass a wide selection of polymers available. The attractiveness of polymers is due to their low cost, variety, solvent, and biocompatibility and tailorable surface properties. Individual components and devices such as microchannel networks, pumps, and valves have been successfully demonstrated using silicon, glass, and polymers. The focus of future research is in three main directions: 1) Development, modification, and optimization of existing microfabrication technologies for low-cost high-volume production of microfluidic devices, 2) Development of new materials and associated microfabrication techniques tailored for specific applications in microfluidics and 3) Development of integrated systems and packaging processes involving multiple materials and fabrication technologies for complex functionality. Combined together, these developments have the potential to radically change biochemical analysis, drug delivery, drug discovery, and tissue engineering. A number of these devices have already permeated into use by non-specialists in microfabrication, demonstrating their long term potential and success.

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Chapter 3 Interfacing Microfluidic Devices with the Macro World

Xuefeng Wang and Kashan A. Shaikh GE Global Research, 1 Research Circle, Niskayuna NY 12309

Keywords: microfluidic interface, chip-to-world, electrospray mass spectrometry, emitter, drug delivery, fluidic couplers

Abstract The universal requirement for all microfluidic devices is a robust fluidic interface between the device and the macro world. This interface consists of two main types of connections: 1) fluid delivery to the device from the macro world, and 2) output from the device to the external surroundings. Microfluidic interfaces must satisfy a number of different criteria including being reliable, mechanically robust, leak free, simple to assemble, and having minimal dead volume. Additionally, since the interface typically requires millimeter sized parts, fabrication processes deviate from the typical MEMS process. This chapter explores the current state of the art in worldto-chip and chip-to-world microfluidic interfaces. World-to-chip interface examples mainly consist of an integrated coupler that mates external capillary tubing to the device. Chip-to-world interface examples include interfacing the microfluidic device to other instruments or directly to biological subjects (e.g., humans).

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3.1 Introduction Microfluidic systems have been widely studied in recent years for chemical detection, biological analysis, medical therapeutics, and pharmaceutical applications. The small volume and large surface to volume ratio enabled by this technology allows chemical and biological processes to be realized with less sample, lower cost, faster speed, and better accuracy than traditional laboratory processes. A microfluidic device typically consists of a combination of fluidic control and sensing components, such as channels, chambers, valves, pumps, mixers, heaters, and sensors. While selective integration of these components is largely application dependent, all devices require an interface between the microdevice and macro world. There are two main types of interconnections: the interface to microfluidic device from macro world for sample loading, and the interface from microdevice to external world for fluidic output. Though numerous passive and active microfluidic components have been demonstrated to date, the microfluidic interface remains an area that is relatively overlooked and under-studied. However, a reliable microfluidic interface is critical for any microfluidic product to be commercially viable.

3.2 Typical Requirements for Microfluidic Interfaces The solution for a successful microfluidic interface must be able to address several issues. First, it needs to be reliable, mechanically robust, and leak free. This allows fluidic samples to be introduced into or out of microfluidic chip securely and without contamination. Secondly, it needs to avoid introducing dead volume. Due to structural mismatches between the interface and microchannel, dead volume is easily created. Existence of dead volumes may reduce the efficiency of the microfluidic system and introduce sample contamination issues. Lastly, it needs to allow simple assembly and the flexibility to attach and detach with relative ease. Many microfluidic devices are designed for disposable use to avoid sample crosscontamination. This requires simple mechanisms to connect devices to external instruments in order to allow for efficient operation.

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3.3 Review of Microfluidic Interfaces 3.3.1 World-to-chip interfaces 3.3.1.1 Direct sample loading structures

The most commonly used interconnection between a microfluidic system and macroscopic world is the loading well [1]. Loading reservoirs, with one end open to the ambient environment and the other end connected to the microchannel network, can be created on various types of substrates using etching or machining techniques [2]. The size of loading reservoirs is typically in the millimeter scale to facilitate manual or automatic liquid loading using syringe needles or pipettes. The liquid loaded in the reservoir can then be drawn into the microchannels by capillary force, differential pressure, or electrokinetic force. 3.3.1.2 Interface adaptors

Interface adaptors are comprised of mechanical parts assembled together and are designed to quickly connect the external world to the microfluidic device. Unlike disposable microfluidic chips or cartridges, the interface adaptors are typical made using conventional machining tools and are intended for repeated use. Advantages include improved mechanical integrity and simplified assembly. Nittis et al. developed a mechanical assembly that provides an interface for coupling standard capillary tubing to microfluidic mixer chips [3]. As shown in Figure 3.1, the assembly measures 38x35x30 mm, and can be assembled in 5 min. When assembled, the recesses in the two main body halves form portholes to position the capillary tubing into the inlets of the microfluidic chip. The alignment of the microfluidic chip and the capillaries is achieved through the alignment guides on the precision positioner that holds the chip. The sealing membrane, which is made from a silicone elastomer, seals off the edges of the inlet and outlet holes to prevent leakage. In addition, the flexible membrane can deform to absorb any machining inaccuracy of the assembly. The assembly also has a glass observation window for optical visualization. In experiments, the interface module withstood flow rates up to 1.5 mL/min before fluid leakage was observed. Since the capillaries enter the chip directly, the dead volume at the fluidic interface is minimized. In addition, the interface does not use any adhesive material, which can cause contamination and clogging issues.

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Fig. 3.1 Schematics of a mechanical assembly microfluidic interface de-veloped by Nittis et al. [3]: (a) assembled module; (b) exploded view (This figure is reproduced by permission of The Royal Society of Chemistry).

Fig. 3.2 Schematics of the microfluidic interface developed by Oh et al. [4]: (a) exploded view; (b) cross-section view of sample loading mode; (c) cross-section view of sample sealing mode (This figure is reproduced by permission of The Royal Society of Chemistry).

Oh et al. developed a microfluidic interfacing system with built-in valves for sample loading and sealing and demonstrated it on a multi-chamber po-


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lymerase chain reaction (PCR) chip [4]. The interface module consists of two plastic fittings and a plastic chip holder. In operation, a PCR chip is first inserted into the chip holder. Then the plastic fittings with rubber sealing sheets are assembled with the holder. PCR samples are loaded into the microchip directly through guide holes in the plastic fittings using pipettes. After the samples are loaded, the plastic fittings with the rubber sheets can be slid to seal the inlets and outlets of the chip without dead volume. Biocompatible Polydimethylsiloxane (PDMS) material is used to make the sealing sheet, to resist pressure building up inside the microfluidic channels. Leakage tests were conducted at elevated temperature (100oC, 30 min) on sealed chips and showed no detectable leakage. Real-time PCR assays were also conducted and yielded a 100% success rate with no contamination or leakage. 3.3.1.3 Micromachined fluidic couplers

Glass or plastic tubing is normally used in chemical analysis systems to transfer fluidic samples. Capillary tubing can be directly connected to a microchip using an adhesive bonding method. To improve the structural integrity, fitting holes can be created on glass or Si substrates using a drilling or etching method. Capillaries can be inserted in the fitting holes and be fixed by adhesives such as epoxy or crystal bond. For capillary electrophoresis applications, glass substrates are preferable since they can sustain the high electrical voltage needed in the tests. For glass chips, Bings et al. reported that fitting holes drilled by standard pointed drills have a conicalshaped bottoms that lead to a dead volume at the capillary and microfluidic system interface [5]. The dead volume is found to result in significant band broadening effect. Bings et al. demonstrated that a flat tipped drill bit could be used to flatten out the conical shaped bottom of the fitting hole to minimize the effective dead volume at the capillary-microchip interface. Li et al. demonstrated use of a PDMS interconnect piece for coupling glass or plastic tubing to microfluidic channels [6]. Holes are punched into a piece of PDMS pad using a glass capillary. The PDMS pad can then be bonded (permanently or reversibly) to microfluidic systems made from Si, glass, PDMS, or other materials. Permanent bonding can be achieved by oxygen plasma treating of bonding interfaces prior to bonding or by using epoxy glues. Glass or Teflon plastic tubing can be inserted into the holes in the PDMS connector. The interconnection can be further enforced by applying epoxy glues. The bonding strengths, leakage rates, and pull-out forces were characterized by Li et al. and it was found that the circularity

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of the holes punched in the PDMS and the cleanliness of the bonding surfaces are critical to achieve high bonding strength.

Fig. 3.3 Schematic of PDMS interconnect developed by Li et al. [6] (This figure is reproduced by permission of the IEEE, ©2003 IEEE).

Due to geometric mismatches, tubing connections are known to introduce dead volumes into the microfluidic system. To alleviate this problem, Chiou et al. used a cast PDMS channel to bridge between a capillary and a microfluidic channel, as shown in Figure 3.4 [7]. When fabricating the fluidic interface, a capillary is first placed close to the opening of a microchannel. A thin metal wire is then threaded into the microchannel through the hollow capillary tubing. PDMS precursor is poured into a junction between the capillary and the microchannel. The PDMS is cured, locking the relative position of the capillary and the microchannel. Then the metal wire is removed from the PDMS, leaving a connection channel bridging between the capillary and microchannel. Due to surface tension, the PDMS connection channel has conical shaped ends with minimized dead volume. The length of the connection channel inside of the microfluidic system can be roughly controlled by pre-curing of the PDMS mixture and by varying the curing temperature. When needed, the capillary can be pulled out and replaced with another one of the same dimension. Leakage tests showed no leakage at the capillary-PDMS interface at pressures below 150 psi or flow rates below 50 µL/min. Pullout tests also showed the mechanical strength of the interface can be adjusted by varying the length of embedded capillary in the PDMS junction, with forces on the order of several Newtons obtainable.


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Fig. 3.4 Schematic of the minimal dead volume microfluidic interconnects from [7] (This figure is reproduced by permission of the IOP).

Han et al. developed a microfluidic system interface using the stereolithography (SLA) technique [8,9]. SLA is an additive fabrication method that uses a laser to fix photosensitive polymer structures layer by layer. It allows complex 3-dimensional microfluidic interfaces to be fabricated with 25 µm vertical (out-of-plane) resolution and 1 µm horizontal (in-plane) pattern resolution. Since the structures are created in a layer-by-layer fashion, integration of external subjects, such as electrical, mechanical, and optical components, into the SLA fabrication process is possible. Shown in Figure 3.5 is a cross-sectional view of the reported microfluidic interconnect and channel system. The one-step plug-in interconnects are made from SLA, with rubber O-rings assembled post fabrication. The interconnect chip can be aligned onto the glass channel chips using tooling holes or alignment marks. A low viscosity epoxy adhesive is then flowed into the gap between the interconnect and the glass chip through a dedicated adhesive via. The epoxy is then cured, to permanently bond the interconnect and the channel chips. To introduce the sample fluids, glass capillary tubing is plugged into the inlet and outlet ports in the interconnect chip. The first rubber O-ring between the interconnect layer and the glass chip is used to prevent adhesive from flowing into and clogging the microchannels. The second O-ring embedded in the interconnect ports is used to provide a tight seal when external capillaries are inserted into the interconnect ports. The pullout pressures of the capillary tubing depend on the diameter of the tubing and the number of O-rings. It was shown that with two Orings, the pullout pressure is about 2 MPa for tubing with inner diameters from 0.25 to 1 mm. Han et al. applied the SLA enabled interfacing technique in a multi-chip genetic sample preparation system [8]. The system consists of several functional components for cell purification, cell separation, cell lysis, solid phase DNA extraction, PCR, and capillary electrophoresis.

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Fig. 3.5 Cross-sectional view of microfluidic interface fabricated using SLA by Han et al. [8] (This figure is reproduced with permission from Elsevier, ©2007, Elsevier).

Gonzalez et al. designed an on-chip fluidic tubing adaptor to interface their microfluidic system with the macro world [10]. The tubing adaptor is essentially a micromachined Si tube, which can connect to a microfluidic channel on one end and external plastic tubing on the other end, as shown in Figure 3.6. The authors fabricated the Si tube on the same chip as the microfluidic channels. Each hexagonal shaped Si channel consists of two halves fabricated simultaneously using anisotropic etching of a (100) silicon wafer on both sides. The two halves are joined together using fusion bonding methods. Plastic tubing with appropriate diameter can then be fitted over the hexagonal tube adaptor. Heat can then be applied at the junction to create a leak-free seal. To provide structural support and mechanical integrity, guide bars can be designed and fabricated on either side of the Si tube adaptor.

Fig. 3.6 Si based tubing interconnect developed by Gonzalez et al. [10] (This figure is reproduced with permission from Elsevier, ©1998, Elsevier).


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Gray et al. used the deep reactive etching method to create fluidic couplers in a silicon substrate for interfacing microfluidic systems with the macroscopic world [11,12]. The accuracy of microfabrication techniques allows high-density fluidic interconnections of arbitrary shapes and sizes to be made with precision. The authors reported three types of fluidic couplers made using this method. A first type of coupler was made by etching circular recess holes into the silicon substrate. The recess holes are connected to the micro channels in a microfluidic system. Since the size of the recess holes matches the diameter of capillary tubing, standard capillaries can be inserted into the recess holes and fixed in place using epoxy adhesives (Figure 3.7a left). Fluids can be transferred into the microfluidic systems through the capillaries using syringe pumps. To avoid possible seeping of the adhesive into the fluidic channel, a second type of fluidic coupler with a circular sleeve structure inside the recess hole was designed. The gap between the outer sleeve wall and the sidewall of the recess hole matches the thickness of the capillary tubing (Figure 3.7a right). When the capillary is plugged into the coupler, the sleeve serves as a barrier to the adhesive and enhances the mechanical robustness of the interface, though introducing some dead volume.

Fig. 3.7 Interconnect coupler structures developed by Gray et al. [11]. (a) Couplers requiring the use of adhesive. Capillary is placed directly in an etched hole (left) or in an etched circular sleeve (right). (b) Coupler using a plastic fitting to hold capillary in place (These figures are reproduced with permission from Elsevier, ©1999, Elsevier).

To eventually circumvent the use of adhesives, a third type of fluidic coupler was developed utilizing plastic press fittings, shown in Figure 3.7b. In this configuration, the capillary tubing is held in place by the plastic fittings. The plastic fitting can be made using injection molding of polyoxymethylene (POM). To integrate the plastic fittings into the system, additional through wafer holes are made using deep reactive ion etching or ultrasonic drilling methods. When the plastic fittings are assembled with the microfluidic chip, plastic pegs are melted with a heat gun to achieve

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permanent assembly. The authors demonstrated the functionality of these microfluidic interfacing methods in a multi-level laminating mixer system. Pan et al. developed an adhesive-free method to form interconnections between microfluidic systems and the external world [13], as shown in Figure 3.8. In this method, a silicon flange is first created, which consists of s through wafer channel surrounded by a concentric sleeve structure. The silicon flange is formed by a two-step etching process including an anisotropic DRIE and an isotropic silicon reactive ion etching (RIE). Only one mask is needed to form the silicon flange and the RIE lag effect is used to differentiate the etch rate at the through wafer channel and the flange sleeve region. To interface with the external world, a silastic tube is connected to the silicon flange using a polyolefin heat shrink tubing. The polyolefin tube has a shrink temperature of 143oC and an expansion ratio of 2:1. When heated above the shrink temperature, the inner diameter of the polyolefin tube shrinks to half of its original size, forming tight seals at both the silicon flange and silastic tube ends. Pullout tests showed that the adhesion between the heat shrink tube and the silicon flange was dependent on the shape of the circular silicon sleeve structure. Among different shapes, circular sleeves showed maximal pullout forces of ~3N, due to large contact area between silicon sleeve and polyolefin tube. Leakage test showed that the heat shrink tubing connection had zero leakage at up to 29 psi for over 24 hours. Moreover, the polyolefin tubing becomes soft when heated to above 180oC and can be removed from the silicon flange, making the silicon microfluidic chip reusable. Since this method does not use any liquid adhesives, clogging of microfluidic channel due to adhesive seepage can be avoided.

Fig. 3.8 Schematic of adhesive-free microfluidic interconnect developed by Pan et al. [13] (This figure is reproduced by permission of the IEEE, ©2006 IEEE).


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3.3.2 Chip-to-world interfaces In many cases microfluidic technology can be useful for simplifying or optimizing a task that involves delivery of fluids to a macro-sized object. Whether this object is an analytical instrument or the human body, microscale systems offer advantages in terms of automation, fluid volume, and added functionality. Efficient connections between the chip and macro world are vital to the success of these microsystems. This section explores two major examples of chip-to-world interfacing: chip-based electrospray for mass spectrometry, and drug delivery/injection. 3.3.2.1 Mass spectrometer interfacing

A very important analytical tool in the life sciences is the mass spectrometer (MS). It can provide accurate determination of a molecule’s molecular weight, requiring only a few thousand molecules. A key parameter in the sensitivity of MS is the efficiency of ionizing the molecules in a sample. Conventional electrospray ionization techniques often form large fluid droplets that require high voltages in order to ionize completely. A major motivation for moving towards the micro-scale is the reduced volume and ability to integrate other pre-MS functionalities (e.g., liquid chromatography or capillary electrophoresis). A smaller sample volume (smaller droplet) allows for a smaller ionization voltage, which then allows the chip’s emitter to be placed closer to the MS ionization inlet. Also, the sensitivity may be increased due to the greater efficiency of ionizing a smaller sample volume. Chip-based MS interface development can be classified into three main areas: (1) electrospray directly from the chip edge, (2) mated capillary emitter, and (3) integrated emitter formed during the microfabrication process [14]. Initial attempts at interfacing microfluidic channels with MS utilized chip edge emitters, since fabrication was fairly straightforward. Karger’s group at Northeastern University was the first to publish results with this concept [15]. They created channels in glass chips via standard photolithographic and wet etching procedures. A total of nine individual channels were built with dimensions of 60-µm wide by 25-µm deep. Each channel extended right to the edge of the glass chip (Figure 3.9). Applying a high voltage between the sample inlet of one channel and the ionization inlet of the MS caused electrospray from the channel outlet. Due to buffer conditions (low electrical resistance or low pH), electroosmotic flow was small and they found it necessary to augment the electrical injection with pressure driven flow. A major issue with the chip edge emitter design is the spreading of

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fluid along the edge of the chip, which is dependent upon the polarity of the fluid and hydrophillicity/hydrophobicity of the edge. This leakage could contaminate neighboring channels and effectively increases the fluid volume at the emitter, counteracting the benefits of moving to the microfluidic scale. If capillary electrophoresis (CE) was integrated on the device, the increased volume at the edge would recombine the separated fractions and render the CE useless. Xue et al. minimized the spreading issue for aqueous solutions by coating the chip edge with a reagent (i.e. Imunopen or n-octyltriacetoxysilane) that made the surface hydrophobic.

Fig. 3.9 Diagram of a system for performing chip edge electrospray. (This figure is reprinted with permission from [15]. Copyright 1997 American Chemical Socety).

Ramsey et al. implemented a chip edge emitter design and additionally integrated an electroosmotic pumping scheme [16]. As shown in Figure 3.10, a side-arm channel was placed very close to the edge emitter and was coated with a linear acrylamide to prevent electroosmotic flow within the channel (increased surface viscosity). By applying a 1.2-kV potential difference between the sample inlet and side-arm channel, electroosmotic flow was induced. The MS ionization inlet was grounded resulting in electrospray from the edge emitter due to a 4.8-kV potential difference between the side-arm channel and MS inlet. Separation between the chip edge and MS inlet was between 3-mm to 5-mm. The chip was scored and physically cleaved to produce the edge emitter. This method could possibly leave a rough edge and lead to locally high electric fields, which would yield a non-uniform electrospray.


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Fig. 3.10 (a) Schematic of a chip edge electrospray emitter with a side-arm channel to facilitate electrosmotic pumping. (b) Photograph of electrospray in action. (These figures are reprinted with permission from [16]. Copyright 1997 American Chemical Society).

Fig. 3.11 PET device for chip edge electrospray. (These figures are reprinted with permission from [17]. Copyright 2001 American Chemical Society).

As shown in Figure 3.11, Rohner et al. used a laser ablation process to fabricate an edge-based electrospray emitter in plastic (PET) [17]. A carbon ink electrode was embedded in the structure for electrospray operation. Since PET is hydrophobic, droplet formation at the chip edge was minimized for aqueous solutions. The use of photoablation and plastic lamination fabrication methods allow for low cost production and one-time-use devices. Plastics are not as optimal for CE separations as glass, so surface modifications may be necessary in order to integrate CE.

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Fig. 3.12 Chip edge electrospray device integrating a nebulizer and CE separation. (This figure is reprinted with permission from [18]. Copyright 1999 American Chemical Society).

Zhang et al. took the initial work from Karger’s group a step further by integrating a nebulizer to minimize the droplet formation at the glass chip’s edge emitter outlet [18]. As shown in Figure 3.12, two auxiliary channels met at the exit of the main channel, constricting the droplet via pressurized flow from those channels. These channels also allowed for buffer modification prior to electrospray. They demonstrated on-chip CE separation and optimized electrospray emission into the MS inlet. By implementing a nebulizer and eliminating droplet formation on the chip edge, CE was achievable. Band broadening caused by serpentine channels and nonoptimal auxiliary channel design resulted in MS efficiencies lower than conventional methods. Design optimizations would likely increase the efficiency. In order to avoid the issues encountered with chip edge emitters, researchers have invented ways to couple glass capillaries to the microfluidic devices. Harrison’s group at University of Alberta drilled into the side of their chip to form a 200-µm diameter cavity into which a fused silica capillary was inserted [5]. They found that drill bits with flat ends could create holes optimal for capillary mating with low dead volume. Using this method they were able to integrate on-chip capillary electrophoresis with subsequent electrospray through the attached capillary and MS analysis (Figure 3.13) [19]. The manual drilling and assembly associated with this method are inherently error-prone and not optimal for mass production, which limits its use for industrial applications.


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Fig. 3.13 CE device mated with a glass capillary for electrospray. (This figure is reprinted with permission from [5]. Copyright 1999 American Chemical Society).

Recent work has focused on developing emitters that are formed during the microfabrication process and overcome the issues of chip edge emitters. Tang et al. demonstrated laser machining in polycarbonate to form a 2D array of nine emitters (Figure 3.14) [20]. These cone-shaped structures avoid large droplet formation and the hydrophobicity of the plastic further prevents liquid spreading. Emission results indicated an increase in MS sensitivity due to the array format since more ions were generated per run.

Fig. 3.14 2D array of cone-shaped emitters. (This figure is reprinted with permission from [20]. Copyright 2001 American Chemical Society).

As shown in Figure 3.15, Kameoka et al. sandwiched a triangular parylene film in between two laminated Zeonor sheets (a polyolefin) [21]. This triangular structure acted as a wick to direct fluid flowing out of microfluidic channels in the Zeonor film. A stable taylor cone was achievable with this device. The Zeonor channels were formed by hot embossing. Manual alignment was necessary to position the parylene tip prior to bonding the

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two Zeonor sheets. Kameoka also demonstrated an array of tips, but each tip had to be separately handled and aligned. The manual nature of the process limits its use as a commercial solution.

Fig. 3.15 Triangular tip emitter sandwiched between two Zeonor sheets containing microfluidic channels. (These figures are reprinted with permission from [21]. Copyright 2002 American Chemical Society).

Yin et al. developed a polyimide-based device integrating a liquid chromatography column, enrichment column, and nanoelectrospray emitter tip [22]. The device was fabricated using laser ablation and is being sold by Agilent for use in its HPLC LC/MS instrument (Figure 3.16). Both the use of polyimide (plastic) and the triangular tip produce optimal conditions for the electrospray operation. The addition of sample enrichment (concentration) and on-chip chromatographic separation greatly facilitates the ability to rapidly perform MS analysis on complex samples.

Fig. 3.16 Polyimide device integrating chromatography, sample enrichment, and electrospray. (This figure is reprinted with permission from [22]. Copyright 2005 American Chemical Society).


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A few groups have focused on developing specialized nozzle emitter tips (Figure 3.17) [23-26]. Of these, the work from Shultz et al. has been successfully commercialized by Advion BioSciences [26]. The nozzle is fabricated in silicon using DRIE and can be as small as 15-µm in diameter. A sample delivery tube interfaces with the back of the chip, transporting the sample directly to the tip. Nozzles allow for minimal droplet formation and can accommodate liquids with varying polarity. Advion developed an array of nozzles and an automated instrument to selectively electrospray through one tip at a time. This design allows each tip to be one-time-use, provides a means to analyze multiple samples using the same chip, and results in a cost-effective, mass-producible commercial product.

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Fig. 3.17 Nozzle emitter tips. (a) [23] (This figure is reproduced by permission of the IOP), (b) [24] (This figure is reproduced by permission of The Royal Society of Chemistry), (c) (This figure is reprinted in part with permission from [25]. Copyright 2003 American Chemical Society), (d) (This figure is reprinted in part with permission from [26]. Copyright 2000 American Chemical Society).

3.3.2.2 Chip-based drug delivery/injection

Microfluidics and microelectromechanical systems (MEMS) are attractive options for drug delivery systems since they are capable of providing precise dosage and spatial control. Much research has focused on developing microneedles for extracting fluids (i.e. blood) from the human body. Compared to conventional needles, microneedles produce much less pain in the patient due to their small size. Delivering fluids using microneedles or other methods has begun to gain interest as well. The major risk associated

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with microneedles is the possibility of breaking the needle during insertion or removal (leaving the needle embedded in the skin). By designing the needle carefully, the risk can be minimized or eliminated. This section presents a few examples of microfluidic coupling to biological tissues for delivery of fluids. Paik et al. created in-plane silicon microneedles integrated with a PDMS microfluidic chip (Figure 3.18) [28]. Fabrication consisted of etching channels in the silicon via DRIE, sealing them by depositing a thick polysilicon layer, and then etching again to form the needle structure. Testing different needle geometries revealed that a 30 degree taper and isosceles triangle shape resulted in optimal resistance to mechanical failure under buckling load and needle insertion. A linear array of needles was shown to be effective at delivering a dye into model systems.

Fig. 3.18 Top: Array of silicon microneedles with buried microchannels. Bottom: The array was used to demonstrate successful delivery of Rhodamine B into a model system [28] (This figure is reproduced with permission from Elsevier, ©2004, Elsevier).


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Fig. 3.19 Silicon microneedle with integrated fluid conduits formed in photosilicate glass [29] (This figure is reproduced by permission of The Royal Society of Chemistry).

Zappe et al. fabricated a silicon microneedle attached to a pyrex wafer with etched microfluidic channels, as shown in Figure 3.19 [29]. Conduits 6.1µm and 2.3µm tall inside the needle were formed with a sacrificial photosilicate glass (PSG) layer. The needle was combined with an automated system for injecting drosophila embryos with RNA interference (RNAi) probes, resulting in a 10-fold efficiency increase compared to manual methods.

Fig. 3.20 Parylene microneedle with gold electrodes for fluid stimulus delivery and neural recording [30] (This figure is reproduced by permission of the IEEE, ©2006 IEEE).

Ziegler et al. used molding and thermal bonding of parylene to create a microfluidic channel integrated with a flexible neural probe [30]. As shown in Figure 3.20, integrated gold electrodes provide neuron recording capabilities for the envisioned drug response analysis system. Fluids could be delivered through the needle to neurons in the brain while measuring neuronal response.

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Blake et al. built a PDMS perfusion chip to accurately deliver stimuli to specific regions of a neonatal rat brain slice (Figure 3.21) [31]. Conventional perfusion systems lack the ability to controllably vary the stimuli across the tissue. By immobilizing the brain slice on microfabricated PDMS posts and flowing solutions through the PDMS channels, the stimuli could be spatially controlled. Electrodes attached to the tissue allowed for real-time monitoring of the response to the stimuli.

Fig. 3.21 PDMS device for perfusion of an immobilized brain slice and stimuli response monitoring [31] (This figure is reproduced by permission of The Royal Society of Chemistry).

3.4. Future Perspectives As indicated by the various examples presented in this chapter, microfluidic interfacing has come a long way. The next step is to move from research level interfaces to more robust, easily manufacturable, and simple to use connections. While there are a few successful examples in industry,


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microfluidic interfacing is often a major hurdle that is not easily overcome. In order for a microfluidic device to move towards commercialization, this perennial problem must be solved. Commercial devices often need to be disposable such that the chip must have the capability to be inserted and removed from an instrument regularly. The interface between the chip and instrument must therefore be straightforward to operate while maintaining a robust connection.

References 1. Fredrickson CK, Fan ZH (2004) Macro-to-micro interfaces for microfluidic devices. Lab on a Chip 4:526-533. 2. Delamarche E, Juncker D, Schmid H (2005) Microfluidics for processing surfaces and miniaturizing biological assays. Advanced Materials 17:2911-2933. 3. Nittis V, Fortt R, Legge CH, de Mello AJ (2001) A high-pressure interconnect for chemical microsystem applications. Lab on a Chip 1:148-152. 4. Oh KW, Park CS, Namkoong K, Kim J, Ock KS, et al. (2005) World-to-chip microfluidic interface with built-in valves for multichamber chip-based PCR assays. Lab on a Chip 5:845-850. 5. Bings NH, Wang C, Skinner CD, Colyer CL, Thibault P, et al. (1999) Microfluidic devises connected to fused-silica capillaries with minimal dead volume. Analytical Chemistry 71:3292-3296. 6. Li SF, Chen SC (2003) Polydimethylsioxane fluidic interconnects for microfluidic systems. IEEE Transactions on Advanced Packaging 26:242-247. 7. Chiou CH, Lee GB (2004) Minimal dead-volume connectors for microfluidics using PDMS casting techniques. Journal of Micromechanics and Microengineering 14:1484-1490. 8. Han KH, Frazier AB (2005) Reliability aspects of packaging and integration technology for microfluidic systems. IEEE Transactions on Device and Materials Reliability 5:452-457. 9. Han KH, McConnell RD, Easley CJ, Bienvenue JM, Ferrance JP, et al. (2007) An active microfluidic system packaging technology. Sensors and Actuators B-Chemical 122:337-346. 10. Gonzalez C, Collins SD, Smith RL (1998) Fluidic interconnects for modular assembly of chemical microsystems. Sensors and Actuators B-Chemical 49:40-45. 11. Gray BL, Jaeggi D, Mourlas NJ, van Drieenhuizen BP, Williams KR, et al. (1999) Novel interconnection technologies for integrated microfluidic systems. Sensors and Actuators a-Physical 77:57-65. 12. Gray BL, Collins SD, Smith RL (2004) Interlocking mechanical and fluidic interconnections for microfluidic circuit boards. Sensors and Actuators aPhysical 112:18-24.

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13. Pan T, Baldi A, Ziaie B (2006) A reworkable adhesive-free interconnection technology for microfluidic systems. Journal of Microelectromechanical Systems 15:267-272. 14. Koster S, Verpoorte E (2007) A decade of microfluidic analysis coupled with electrospray mass spectrometry: An overview. Lab on a Chip 7:1394-1412. 15. Xue QF, Foret F, Dunayevskiy YM, Zavracky PM, McGruer NE, et al. (1997) Multichannel microchip electrospray mass spectrometry. Analytical Chemistry 69:426-430. 16. Ramsey RS, Ramsey JM (1997) Generating electrospray from microchip devices using electroosmotic pumping. Analytical Chemistry 69:1174-1178. 17. Rohner TC, Rossier JS, Girault HH (2001) Polymer microspray with an integrated thick-film microelectrode. Analytical Chemistry 73:5353-5357. 18. Zhang B, Liu H, Karger BL, Foret F (1999) Microfabricated devices for capillary electrophoresis-electrospray mass spectrometry. Analytical Chemistry 71:3258-3264. 19. Li JJ, Thibault P, Bings NH, Skinner CD, Wang C, et al. (1999) Integration of microfabricated devices to capillary electrophoresis-electrospray mass spectrometry using a low dead volume connection: Application to rapid analyses of proteolytic digests. Analytical Chemistry 71:3036-3045. 20. Tang KQ, Lin YH, Matson DW, Kim T, Smith RD (2001) Generation of multiple electrosprays using microfabricated emitter arrays for improved mass spectrometric sensitivity. Analytical Chemistry 73:1658-1663. 21. Kameoka J, Orth R, Ilic B, Czaplewski D, Wachs T, et al. (2002) An electrospray ionization source for integration with microfluidics. Analytical Chemistry 74:5897-5901. 22. Yin NF, Killeen K, Brennen R, Sobek D, Werlich M, et al. (2005) Microfluidic chip for peptide analysis with an integrated HPLC column, sample enrichment column, and nanoelectrospray tip. Analytical Chemistry 77:527-533. 23. Griss P, Melin J, Sjodahl J, Roeraade J, Stemme G (2002) Development of micromachined hollow tips for protein analysis based on nanoelectrospray ionization mass spectrometry. Journal of Micromechanics and Microengineering 12:682-687. 24. Schilling M, Nigge W, Rudzinski A, Neyer A, Hergenroder R (2004) A new on-chip ESI nozzle for coupling of MS with microfluidic devices. Lab on a Chip 4:220-224. 25. Svedberg M, Pettersson A, Nilsson S, Bergquist J, Nyholm L, et al. (2003) Sheathless electrospray from polymer microchips. Analytical Chemistry 75:3934-3940. 26. Schultz GA, Corso TN, Prosser SJ, Zhang S (2000) A fully integrated monolithic microchip electrospray device for mass spectrometry. Analytical Chemistry 72:4058-4063. 27. NanoMate Fact Sheet. Advion BioSciences. (2008). 28. Paik SJ, Byun A, Lim JM, Park Y, Lee A, et al. (2004) In-plane single-crystalsilicon microneedles for minimally invasive microfluid systems. Sensors and Actuators a-Physical 114:276-284.


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29. Zappe S, Fish M, Scott MP, Solgaard O (2006) Automated MEMS-based Drosophila embryo injection system for high-throughput RNAi screens. Lab on a Chip 6:1012-1019. 30. Ziegler D, Suzuki T, Takeuchi S (2006) Fabrication of flexible neural probes with built-in microfluidic channels by thermal bonding of Parylene. Journal of Microelectromechanical Systems 15:1477-1482. 31. Blake AJ, Pearce TM, Rao NS, Johnson SM, Williams JC (2007) Multilayer PDMS microfluidic chamber for controlling brain slice microenvironment. Lab on a Chip 7:842-849.

Chapter 4 Genetic Analysis in Miniaturized Electrophoresis Systems

Li Zhu1 and Hong Wang2 1

GE Global Research, 1 Research Circle, Niskayuna NY 12309

2

Department of Chemistry, Louisiana State University, Baton Rouge, LA 70803

Correspondence should be addressed to: Li Zhu ([email protected]) Hong Wang ([email protected])

Keywords: microfluidics, microchip, electrophoresis, miniaturized electrophoresis system, genetic analysis, DNA analysis

Abstract As electrophoretic separation of DNA is one of the most important steps in many genetic analyses, continual advances in the development of miniaturized electrophoresis systems are critical to meet the growing need for highquality and low-cost genetic analyses. The commercialization of these miniaturized systems is having or will have significant impact on the revolutionary research in biomedical and life sciences areas. This chapter focuses on the microchip electrophoresis and its applications in genetic

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analyses. The recent developments in the fabrication of electrophoresis microchips, the sieving matrices, the novel micro-fabricated structures for DNA separation, and the internal wall coating chemistries are reviewed. A survey of recent progress in multiplexing and integrating steps toward a high-throughput, low-cost, and miniaturized electrophoresis system is provided. Some commercial microfluidic instruments for genetic analysis as well as their technologies are discussed and we conclude the chapter with a future perspective of the microfluidic technology in industry.

4.1 Introduction 4.1.1 Status of genetic analyses The Human Genome Project, with the primary goals of identifying all of the approximate 30,000-40,000 genes and determining the primary structure of the entire human genome comprised of its 3 billion base pairs, was initiated in 1990 and completed in 2003, the 50th anniversary of the discovery of the DNA double-helix structure [1, 2]. The successful completion of the Human Genome Project marked a significant milestone in the history of science. However, it does not signify an end to the further pursuit of novel sequencing and genotyping technologies. To the contrary, the demand for genomic sequence information, both in basic biomedical research and in routine clinical healthcare, has never been greater. This has stimulated a new round of competition in innovating high-throughput and cost-effective sequencing and genotyping strategies. In 2005, the National Human Genome Research Institute sponsored several research projects with the near term goal of cutting the sequencing cost of a mammalian sized genome to $100,000, and the final goal of a $1000 genome. DNA sequencing in a microchip by the Sanger method is one of the most promising approaches to achieve the near-term goal [3-6]. Currently, the cost of sequencing a mammalian sized genome with the state-of-the-art capillary array sequencer is around $1 million, which is still cost-prohibitive for most institutions and researchers. As alternatives, a variety of genetic analysis technologies targeting the variations in the genome have been explored. Instead of sequencing every single base along the genome, these technologies only interrogate the sites of interest, which have already been identified as the biomarkers for a specific disease or a specific application. DNA microarrays and electrophoretic separations are

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two complementary methods and the most important approaches in genetic analysis. DNA microarrays, first developed in mid-1990s [7-12], turned out to be a powerful tool for high throughput genetic analysis. They interrogate hundreds to thousands of variations among a number of samples simultaneously. Microarray technology, however, will not be covered in the scope of this chapter. A few recent reviews on microarray technology can be recommended to readers for a quick glimpse of the state-of-the-art of the technology [13-21]. Complementary to microarrays, the size and/or conformation based electrophoretic separations of DNA generally target only one or a few variations in several to hundreds of samples. They are widely used in clinical diagnostics, forensic applications and biomedical research [22-29]. The last two decades have witnessed the transformation of the electrophoresis platform from the slab-gel to the capillaries/capillary arrays and to the present microchip format. This chapter will focus on the recent technology innovations in microchip/microfluidic electrophoresis for genetic analysis. 4.1.2 Genetic analysis by miniaturized electrophoresis system 4.1.2.1 Sanger sequencing of DNA

The Sanger sequencing method has served as the cornerstone for most genome sequencing efforts since first demonstrated by Frederic Sanger in 1977 [30]. It is a polymerase enzyme based replication method, similar to that in PCR, but using some specially designed 2’, 3’-dideoxynucleotides (ddNTPs) instead of all deoxyribonucleosides (dNTPs). The ddNTP terminate the extension reaction at a specific base when they are incorporated into the DNA chain due to the lack of an OH group on the 3’ –carbon position of the deoxyribose sugar. The termination occurs randomly at different positions in different copies, producing a nested set of DNA fragments of different length that are complementary to the unknown sequence (Fig. 4.1). After removing the excess ddNTPs and buffer components in the extension mixture by a purification step, the DNA ladders are electrophoretically separated. By correlating the length of the fragments with the identity of the terminating base through a fluorescently labeled dye, one can determine the nucleotide sequence using electrophoresis. Although various emerging sequencing technologies have been explored for ultra-low cost sequencing, the Sanger sequencing method remains the only viable technology for large genome de novo sequencing [3-5]. Tremendous efforts have been devoted to reducing the reagent consumption

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and increasing the throughput by developing multiplexed and/or integrated electrophoresis microchips in Sanger sequencing. Recently, a read length from a glass electrophoresis microchip was reported comparable to that of conventional capillary arrays, while the throughput was several folds higher and the reagent consumption was one to two orders of magnitude less [31]. Disposable plastic microchips were also tested for Sanger sequencing. However, it is still challenging to achieve the required single base resolution and long read length on those microchips [32, 33].

Fig. 4.1 Top: Chemical strudctures of deoxyribnucleoside (dNTP) triphosphate and dideoxyribonucleoside (ddNTP) triphosphate. Bottom: Schematic diagram of Sanger chain termination DNA sequencing reaction.

Genetic Analysis in Miniaturized Electrophoresis Systems 121 4.1.2.2 Genotyping: polymorphisms and mutations

More than 99% of human DNA sequences are identical across the population. However, the genetic or DNA sequence variations in the genome can have a major impact on predisposition to diseases, response to environmental factors, and the effectiveness of the medicines and vaccines [3436]. When the variations in the genome are present in at least 1% of the population, they are generally not harmful and are called polymorphisms. Otherwise, when the variations are less common (less than 1%) and frequently result in diseases, they are called mutations. A variety of size and/or conformation based electrophoretic separation strategies have been developed for identifying polymorphisms and mutations in the human genome during the past decades [28, 37]. Single nucleotide polymorphism (SNP), a single base substitution, deletion or insertion, is the most abundant variation in the human genome. There are more than 3 million SNPs that have been identified, which corresponds to about 1 in every 1000 bases. Several electrophoretic separation strategies have been used to identify the SNPs and single base mutations, including: restriction fragment length polymorphism (RFLP), single strand conformational polymorphism (SSCP), and heteroduplex analysis (HA). RFLPs are applied when the genetic variations create or delete the sites that can be recognized by specific restriction enzymes. The difference in homologous DNA sequences can be detected by the presence of the differently sized fragments after the restriction endonuclease digestion and the electrophoretic separation. RFLPs can be used in paternity testing and disease diagnosis. Guttman et al. developed a rapid PCR-RFLP method to analyze the mitochondrial DNA mutation in diabetes [38]. Microchip electrophoresis was used by Qin et al. to genotype the -6A/G polymorphism in the core promoter region of the AGT gene using the PCR-RFLP method [39]. SSCP is a simple and versatile method of electrophoretically separating the single-stranded DNA (ssDNA) based on subtle differences in sequence (often a single base pair). While the mobility of double-stranded DNA (dsDNA) depends largely upon the fragment sizes but little on the fragment composition, the mobility of ssDNA can be dramatically influenced by the nucleotide composition of a fragment with a specific three dimensional conformation resulting from the intra-strand base pairing. Tian et al. used microchip electrophoresis to detect the common mutations in the BRCA1 and BRCA2 gene [40]. A 384-lane electrophoresis microchip was tested in genotyping 21 single-nucleotide polymorphisms (SNPs) from

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HFE, MYL2, MYL3, and MYH7 genes associated with hereditary hemochromatosis (HHC) and hereditary hypertrophic cardiomyopathy (HCM) [41]. The HA method is based on the mobility difference between the heteroduplex and the corresponding homoduplex. It is a simple and convenient method that has been widely used in biomedical research and clinical diagnostics [42]. In addition, HA can be combined with SSCP and allelespecific amplification method for rapid identification of mutations [43-46]. 4.1.2.3 Short tandem repeats analyses

Short tandem repeats (STRs) or microsatellites are another common polymorphism predominantly used in forensic identification and paternity testing [23, 47, 48]. STRs are 7 to 20 repeats of specific DNA sequences ranging from 2 to 10 bases long. Recently, a 96-channel microfabricated capillary array electrophoresis device was evaluated for forensic STR typing using commercially available forensic kits, e.g., PowerPlex 16 and AmpFlSTR Profiler [49]. In most cases, only 2-3 base-pair resolution is required for STR analysis, therefore plastic microchips are well-suited for such an application. Shi et al. reported a successful two-color sizing analysis of four-locus (CSF1PO, TPOX, TH01, vWA) STRs in poly (cyclo olefin) microchips [33, 50].

4.2 Microchip Electrophoresis for Genomic Analysis Microchip electrophoresis shares basic principles with slab-gel and capillary electrophoresis, therefore, the DNA migration models, the gel matrices, the surface coating strategies and the fluorescent dyes developed for slab-gel and capillary electrophoresis systems are suitable for microchip electrophoresis as well. Microchip electrophoresis also has its unique properties and advantages, such as its ultra-fast analysis time, ultra-low sample consumption, capability of easy scale-up for high throughput analysis and potential for convenient system automation and integration. More recently, various thermoplastic materials have been explored for the mass production of disposable electrophoresis microchips, making lowcost microchip-based instruments more practical.


4.2.1 Material and fabrication of electrophoresis microchips 4.2.1.1 Glass, quartz, and silicon

Glass, quartz, and silicon are the most widely used materials in microchip electrophoresis due to their high-quality optical properties, welldocumented surface chemistries and well-developed fabrication procedures adopted from the microelectronic industry [51-57]. These microchips are mainly manufactured by standard photolithographic procedures followed by chemical etching. Unfortunately, the fabrication of these glass or quartz-based micro-devices is relatively expensive and time-consuming; which make them less desirable for low cost applications. In addition, isotropicity of the wet etching in conventional fabrication produces shallow, elliptical shaped structures with low aspect ratios. Although high aspect ratio structure can be made by a dry etching method, such as DRIE (dry reactive ion etching), the high cost and long fabrication time are still costprohibitive. Other disadvantages associated with microchips made from these materials include the requirement of high temperatures (600 °C) and the use of harmful chemicals (e.g., hydrofluoric acid). 4.2.1.2 Polymer/plastic materials

To circumvent the limitations associated with glass-based devices, several polymer materials have been explored as the microchip substrates. Polymers offer a wide range of mechanical and thermal properties, providing various selections for different applications and allowing rapid inexpensive mass production of micro-devices. They make the economical single-use devices become possible, eliminating cleaning and sample-to-sample carryover contamination. PDMS (polydimethylsiloxane) is the most popular elastomer used for microchip fabrication [58, 59]. It is an inexpensive, flexible and chemically inert material with optically transparency down to 230 nm and little autofluorescence. The PDMS microchannels can be fabricated by softlithography [59]. Briefly, a negative mold is fabricated by conventional micromachining technology. For example, the negative thick photoresist SU-8 has been widely used to fabricate negative mold through photolithography. Then the PDMS prepolymer (e.g., the 10:1 mixture of base and curing agent) is casted over the mold. After curing, which typically takes a few hours, the PDMS film carrying the microstructure can be peeled off from the mold. This PDMS film can be reversibly sealed to another PDMS film or other ultra-flat surfaces by conformal contact. It can also be irre-

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versibly sealed to another PDMS film or a glass plate after an oxygen plasma treatment. Because of the elastomeric nature, PDMS deforms easily. An interesting and effective approach to overcome this is to fabricate PDMS-glass or PDMS-silicon hybrid microchips, which not only takes advantage of the easy and fast fabrication of PDMS, but also the strength from the rigid glass or silicon materials [60-62]. Thermoplastics are attractive alternatives for mass production of the low cost and disposable microchips as well. The thermoplastics used in electrophoresis microchips include PMMA (polymethylmethacrylate), COC (cyclic olefin copolymers, commercialized as Topas® by Ticona and Zeonex® by Zeon), and PC (polycarbonate). One of the most attractive features associated with thermoplastic materials is their flexibility in the micromachining process [63-67]. Thermoplastic fabrication techniques can be broadly classified into two categories, direct fabrication methods and replication methods. Direct methods, by which the individual polymer surface is fabricated to form microstructures, are mostly used for rapid prototyping. Examples include mechanical milling and laser ablation [68]. They are not the most economical and in some cases produce rough surfaces that are problematic for highresolution electrophoretic separations. Replication methods employ a precise template or molding tool from which identical features can be replicated. Examples are injection molding [69], hot-embossing [70], and imprinting [71]. Such techniques are proven to be cost-effective when large numbers of identical plastic parts or devices are formed. Hot-embossing is an easy and inexpensive method for reproduction of microstructures. Under vacuum conditions, a metal molding tool is pressed into a heated polymer substrate, transferring the desired features into the polymer. The number of daughter parts that can be fabricated from a single master and the average replication time per device is ~5 min. Injection molding is the industrial standard process for manufacturing macroscopic plastic parts, and it has been optimized for microstructure fabrication. The molten thermoplastic particles are transported to the heated mold cavity, which contains negative mold. Then the cavity will be cooled to release the microchip. By adjusting the process time and temperature, injection molding can be used to fabricate 3-D micro-structures with excellent precision. However, the high temperature and pressure during the fabrication process introduce thermal stresses, which make the microchip sensitive to the temperature and pressure fluctuations. Further material and fabrication methods for microchips can also be found in Chapter 2.


4.2.2 Theory of gel electrophoresis of DNA The electrophoretic mobility of DNA in free solution is independent of its size due to the constant charge-to-size ratio. In order to achieve a size separation of DNA, it is necessary to introduce certain physical obstacles on a molecular level. One of the widely used physical obstacles is called sieving matrix or gel, which will selectively retard the migration of DNA molecules under an electric field based on their sizes and conformations. Numerous polymers or gels have been utilized as the sieving matrices in capillary and microchip electrophoresis. Understanding the interaction between the polymer networks and DNA species during electrophoretic separation is critical in optimizing sieving matrices and improving performance of electrophoresis [72-75]. Two widely accepted models of DNA electrophoresis in a gel polymer are the Ogston model and the reptation model. The Ogston model [76] treats the polymer network as a molecular sieve. It assumes that the matrix consists of a long, inert and randomly distributed network of interconnected pores with a certain average pore size, through which the solute molecules migrate as a spherical coil. These unperturbed spherical migrating species diffuse laterally until they encounter pores large enough to accommodate passage. Small molecules migrate faster because they have accessibility to a larger fraction of the available pores. The Ogston model predicts a linear relationship between the log µ, where µ is the electrophoretic mobility, and the gel concentration for small DNA molecules and low electronic field. However, according to Ogston model, the mobility of migrating species will quickly approach zero when the molecular radius approaches the pore size of the sieving matrix. In other words, this model cannot explain the behavior of large, flexible molecules migrating through sieving media with mesh size significantly smaller than their size. This phenomenon can be described by the reptation model. The reptation theory [77] suggests that large DNA molecules, when they are too large to fit through a pore while maintaining a coiled conformation, would exhibit “snake-like” migratory behavior moving through the smaller pore network of the gel under the influence of an electric field. This model is based upon the assumption that the migrating molecule can deform and stretch, behaving as a free draining coil instead of an immutable sphere with fixed radius of gyration described by the Ogston model. The mobility of the analyte molecule by this process is inversely proportional to its molecular size. When the electric field strength increases, reptation turns into a biased reptation model [78]. Under this model, the electric field-induced

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orientation extends the stretching periods of DNA, causing their random walk to become strongly biased in the forward direction. The leading end of DNA becomes completely aligned with the field so that no further field effects exist. In a fully biased reptation regime, the mobility increases to saturation at which point there is no dependence of mobility on DNA length and this results in all large fragments migrating at the same rate. With the fast growth of capillary electrophoresis in 1990s, the aforementioned two models derived from slab-gel electrophoresis have become less convincing when it comes to explaining the high sieving power a linear polymer possesses. There have been inconsistencies between theory and experimental observations. Barron et al. [79] showed that the separation of 2.0-23.1 Kbp DNA fragments is possible in 0.0006% (w/w) uncrosslinked hydroxyethyl cellulose (HEC) polymers. To explain the DNA separation in ultra-diluted and unentangled polymer solution, a transient entanglement coupling mechanism was proposed, which postulated that the DNA fragments hook and then drag the uncharged polymer chains during the migration. The larger DNA molecules have higher chance of encountering and entangling polymer molecules, therefore, experience a greater reduction in mobility. This mechanism also applies to DNA separation in capillary electrophoresis using linear entangled polymer solutions at high concentration. 4.2.3 Gel matrices The DNA migration behavior and the separation efficiency are largely affected by the sieving matrices. Therefore, the choice of the sieving matrices is critical to the success of the separation of DNA. Most of the gel matrices that are applied to capillaries can be applied to microchips as well, if they can be loaded into the microchannels. Compared with their conventional counterpart, microchips cannot sustain as much pressure as the capillary during the matrix loading, e.g. 50 psi for most plastic microchips and 200 psi for glass or silicon microchips. An ideal sieving matrix for DNA separation on a microchip should include the following properties: low viscosity, high molecular sieving power, dynamic coating ability, and cost effectiveness. Recent efforts to improve microchip DNA separations have led to the development of new sieving solutions that are more compatible with the microlfuidic format [80-83]. One example is the development of the thermoresponsive polymer [81]. It possesses lower viscosity for easy loading, and its viscosity changes with temperature to be suitable for separation. However, the optimal performance of such a polymer has not been


fully developed yet. The effort in designing and synthesizing an ideal polymer for DNA separation on a microchip is still continuing. In this section, the traditional gel matrices as well as some novel polymer matrices will be introduced. 4.2.3.1 Cross-linked polyacrylamide gel

Cross-linked polyacrylamide is one of the most commonly used separation matrices for slab gel DNA sequencing. It is prepared by a radical copolymerization of acrylamide, with a varying amount of N, N’-methylene bisacrylamide, a cross-linker. The polymerization is typically initiated by ammonium persulfate (APS) and catalyzed by N, N, N’, N’tetramethylethylenediamine (TEMED). Upon polymerization, a dense, cross-linked and flexible polymer network is formed, which has pore sizes ranging from a few nanometers to tens of nanometers. This tight pore network is effective for high-resolution separations of DNA ranging from 6 bases to about 1000 bases long, depending on the gel concentration and the level of cross-linking. Although having been long standing with the slab gel technology, cross-linked polyacrylamide is rarely used in capillaries or microchips. This is because of a few drawbacks associated with it, such as the gel shrinkage and the formation of bubbles during polymerization, as well as the degradation of the polyacrylamide by alkali hydrolysis during electrophoresis. 4.2.3.2 Polysaccharide and its derivatives

Polysaccharides and their derivatives have been used as sieving matrices for DNA separation in both capillary and microchip electrophoresis [83]. They are usually selected when the high-resolution separation is not required. Low viscosity cellulose derivatives, methylcellulose (MC), hydroxyethylcellulose (HEC), hydroxypropyl cellulose (HPC), and hydroxypropylmethylcellulose (HPMC) were used on PMMA microchips for ultra fast dsDNA separations. Agarose, the most widely used lowresolution slab-gel material, was used in a PDMS-PMMA hybrid microdevice, a glass-silicon, and a PDMS-glass hybrid microchip for double strand DNA separation [84-86]. 4.2.3.3 Linear polyacrylamide (LPA) and acrylamide derivatives

Since the first introduction of replaceable, ultra-high-molar-mass Linear Polyacrylamide (LPA) matrices by Karger et al. [87-90], LPA has been the matrix of choice for high performance DNA separation, and in particular,

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DNA sequencing. LPA gels can be formed by polymerizing acrylamide the same way as cross-linked polyacrylamide, except that no cross-linking agent is added. LPA possesses a few highly desirable properties for DNA separation, particularly its high hydrophilicity and excellent sieving power owing to the physical entanglements of the polymer chains. In addition, these linear polymers are not as rigid as their cross-linked counterpart. Therefore, they can be easily replaced in the capillary or microchip after each run. The first DNA sequencing by CE with replaceable linear polyacrylamide reported a read length of 350 bases from an M13 template in 30 minutes [88]. Later, Karger and co-workers reported an improved sequencing read length of 1300 bases in 2 hours [90] by using an LPA mixture composed of 0.5% w/w 270 KDa and 2% w/w 17 MDa based on their previous result of 1000 bases in less than 1 hour [87, 89]. Although numerous types of water soluble, high-molar-mass polymers have been investigated for sequencing applications, LPA remains the most widely accepted sieving matrix for high-performance DNA sequencing by both capillary and microchip electrophoresis. Despite the superior performance of LPA gels, their high viscosity and lack of intrinsic wall-coating ability make them less suitable for microchips. Their high viscosity makes them difficult to fill into and be removed from the microchannel, and in many cases, requires the application of high pressure. In addition, a rather complex coating procedure is usually required on the microchannel walls prior to electrophoresis to minimize the electroosmotic flow and the analytewall interactions. Barron and co-workers created the concept of “nanogel” by incorporating a low percentage of cross-linker in high-molar-mass LPA [91, 92]. The small amount of cross-linker localizes the cross-linking and keeps the flowing ability of LPA. It has been reported that the nanogel provides 18% longer read length in sequencing than a matched LPA counterpart in a microdevice. Poly-N, N-dimethylacrylamide (PDMA) is a widely used acrylamide derivative for DNA separations. This polymer is commercialized by Applied Biosystems as POP (Performance Optimized Polymer) gels. Advantages of PDMA over LPA are its resistance to hydrolysis, self-coating ability, and significantly lower viscosity. However, the performance of PDMA has been less satisfactory compared with that of LPA. This may in part due to the hydrophobic interaction between the PDMA and the DNA labeling dyes. The best performance achieved was 800 bases with a final resolution of 0.5 in 96 min, and 1000 bases with a final resolution of 0.3 in a capillary [93]. PDMA has been the matrix of choice for a few commercial micro-


chip electrophoresis instruments, such as the 2100 Bioanalyzer by Agilent (www.agilent.com), for which high-resolution DNA sizing (e.g., singlebase resolution) is not required. 4.2.3.4 Thermoresponsive polymer matrices

Thermoresponsive polymer matrices exhibit low viscosity at one temperature for gel loading and high viscosity at another temperature for high resolution separation [81, 82]. They can be divided into two categories: thermo-thickening (thermo associating) polymers and thermo-thinning polymers. Thermo-thickening or thermo associating polymers have low viscosity at the low temperature. The viscosity of the polymer increases when it is heated to the high critical solution temperature (HCST). Pluronic polyol F127 (PEO99PPO69PEO99) [94], a typical block copolymer with thermal self-association properties, was tested for dsDNA separation in a PMMA microchip. A grafted copolymer with a hydrophilic LPA backbone and comb-like pNIPA side chains, it was investigated for DNA sequencing. It delivered read length of 800 bases at the optimized composition [95]. Thermo-thinning polymers, on the other hand, have high viscosity at the room temperature. When heated up, the polymer becomes less viscous. Thermo-thinning random copolymers composed of different ratios of DEA (N,N-diethylacrylamide) and DMA (N,N-dimethylacrylamide) were used for DNA sequencing, in both capillary and microchip [81]. Thermoresponsive polymers are promising alternatives as sieving matrices for microchip electrophoresis; however, their performance still needs to be optimized. It would be difficult for these polymers to beat the superior performance of LPA, mainly due to the hydrophobic moieties required for the thermo-response. 4.2.3.5 Other Polymers: PEO, PVP, and PVA

Other replaceable polymer solutions, such as polyethylene oxide (PEO), polyvinyl pyrrolidone (PVP), and polyvinyl alcohol (PVA), have been developed and reported for both DNA fragment analysis and sequencing. PEO is a self-coating polymer widely used in DNA separation. It requires a surface pre-treatment as long as 2-hours with HCl to enhance the silica surface absorption before electrophoresis. Yeung’s group achieved a read length of up to 1000-base with PEO at the optimized conditions in a capillary [96, 97]. Xu et al. systematically investigated the performance of PEO

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in separating DNA ladders at various combinations of molecular weight and concentrations in PMMA microchips [98]. PVP is another polymer with self-coating ability but also a low viscosity. It can provide moderate separation efficiencies under optimum gel compositions. Several groups have demonstrated DNA fragment sizing in both capillaries and microchips using such a polymer [99-102]. The main obstacle for the routine use of PVP in DNA separation is the large mobility shifts of fluorescently labeled DNA fragments observed in this matrix. PVA is resistant to hydrolysis over all pH ranges, and it maintains very low viscosity even at concentrations up to 12%. It was reported as a sieving matrix for fragment analysis in a capillary [103]. However, its separation efficiency diminishes very quickly due to the strong hydrogen bonding presented in the polymer, resulting in self-aggregation. PVA can also be used as a coating polymer to reduce the EOF and analyte absorption onto the walls. 4.2.4 Novel DNA separation strategies on microchips Recently, alternative mechanisms for DNA separation without using a sieving matrix have been introduced for microchip electrophoresis. One approach is to label the DNA fragments with large uncharged molecules, e.g. streptavidin or long chain PEG, to break the constant charge to size ratio. The longer end-labeled DNA fragments migrate faster than the shorter ones, so the size dependent separation can be achieved in a free solution. This method is called end-labeled free-solution electrophoresis (ELFSE) [104, 105]. A number of novel separation strategies employing microfabricated mechanical obstacles at a size scale comparable to the radius of DNA gyration, including micro- and nano-pillar arrays, nano-spheres, and nano-sized channels, have also been developed. The idea of replacing the gel or polymer sieving matrices with micrometer or sub-micrometer sized pillar arrays was first demonstrated by Volkmuth and Austin, who fabricated a rectangular array composed of 1 µm diameter cylindrical posts with 2 µm center-to-center distance on a silicon wafer by optical microlithography [106, 107]. Comparable to the nominal pore size of a 0.05% agarose gel, this array could resolve the DNA fragments up to 100 Kbp. Similar arrays were also employed for pulsed-field electrophore-


sis [108] and continuous macromolecule sorting [109] by the same group. A much more complex and powerful demonstration of this array format was a “DNA prism” for high-speed continuous sorting of DNA fragments in different directions according to their sizes, just as an optical prism deflects the light at different angles according to the wavelength (Fig. 4.2) [110]. Under a symmetric pulsed field, the DNA prism could separate 61209 Kbp DNA molecules in 15 seconds with ~13% resolution, which is 1000 times faster than the conventional pulsed field gel electrophoresis (PFGE) (10-240 hours), and 40 time faster than the pulsed field capillary electrophoresis (~40 minutes). To improve the performance of small DNA fragment separation, Kaji et al. fabricated sub-micrometer, high aspect ratio nano-pillars (100-500 nm diameter and 500-500 nm tall) to resolve DNA fragments ranging from 1 to 38 Kbp within a 380 µm-long channel in 10 seconds [111].

Fig. 4.2 Structure of the microfabricated device illustrating the sieving matrix integrated with the microlfuidic channels. The post array is 3 mm by 9 mm, and the posts are 2 µm in diameter, 2 µm apart, and 2 µm tall. A single channel connecting to the DNA reservoir injects DNA through a 28-V opening. The many microfluidic channels connected to buffer reservoirs produce uniform electric fields over the sieving matrix by acting as electric-current injectors. (Reprinted by permission from Macmillan Publishers Ltd: Nature Biotechnology [110], copyright 2002.)

In contrast to the pillar arrays fabricated by conventional photolithography technology, Doyle et al. introduced a self-assembled, quasi-regular temporary pillar array by applying a constant and homogeneous magnetic field to a suspension of paramagnetic particles confined in a microfluidic channel. By alternating the size and the concentration of the magnetic particles, the

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nominal “pore” size of the array can be altered for different applications [112]. A separation of λ-phage, 2λ-DNA, and bacteriophage T4 DNA was achieved in as short as 150 s and resolutions greater than 2 between λ and T4 were successfully obtained [113]. In addition, the movement of a single T4 DNA molecule (169 Kbp) in the magnetic particle pillar array and a theoretical model were presented [114]. Brownian motion has been exploited by several groups for the size dependent DNA separation. One approach is to fabricate an asymmetric array of obstacles as a Brownian ratchet to rectify the Brownian motion laterally and thereby deflect diffusing particles depending on their size. In a 4-inch (10-cm) silicon wafer, a nominal 6% resolution by length of DNA molecules in the size range of 15–30 Kbp was demonstrated by Chou el al. in 1999 [115], followed by a continuous DNA sorting device by Cobadi et al. in 2002 [116]. To speed up the separation based on the intrinsically slow Brownian motion, a tilted electric field was applied to the array [117]. The time required for resolving 48.5 Kbp and 164 Kbp DNA fragments was reduced from 140 min to 14 min, a 10-fold improvement. Alternatively, a Brownian ratchet could be created by the electric field instead of physical obstacles, as demonstrated by Bader et al [118, 119]. More recently, another Brownian motion based phenomenon, absolution negative mobility (ANM), was explored for separating DNA fragments [120, 121]. Another intriguing approach is the entropic based separation, which was demonstrated by an “entropic trap” device [122] (Fig. 4.3) and an “entropic recoil” device [123, 124] (Fig. 4.4). The basic principle of entropic based separation is that DNA molecules are energetically favorable to be in the coiled state since this maximizes their entropy. The entropic trap device is a microfluidics channel defined by a series of alternating thin (75100 nm) and thick (1.5-3 µm) regions in the microchannel. The DNA molecule can keep in the coiled state in the thick region; however, when it meets the thin region where the channel size is much smaller than the gyration of a DNA molecule, the DNA has to deform to pass through. Since the deformation is entropic unfavorable, the DNA molecule is temporarily trapped at the entrance of the thin region. An escape of the whole DNA molecule is not initiated until majority of that molecule is introduced into the thin region by Brownian motion. Interestingly, the longer molecules have a larger contact area and consequently a higher probability to escape the entropic trap, therefore, they move faster than the shorter ones. The entropic recoil device consists of a high entropy pillar-free planar region and a low entropy pillar-filled region on a chip. After applied the electric field, the DNA molecules are forced to move to the pillar region. When the elec-


tric field is removed, the molecules will recoil to the planar region unless they have entirely moved to the pillar region. In contrast to the entropic trap, the shorter DNA molecules move faster than the longer ones. A rapid separation of T2 (167 Kbp) and T7 (39 Kbp) was demonstrated. Applying a voltage pulse of 2 s, the shorter T7 molecules were fully inserted, while the longer T2 molecules remained partially in pillar-free region and subsequently recoiled.

Fig. 4.3 Nanofluidic separation device with many entropic traps. (A) Crosssectional schematic diagram of the device. (B) Top view of the device in operation. (C) Experimental setup. (From [122]. Reprinted with permission from AAAS.)

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(a)

(b)

Fig. 4.4 (a) DNA molecules are driven into the dense-pillar region, which occupies the bottom 80% of each frame. (b) Entropic recoil at various stages ending when all the molecules have recoiled except the leftmost. (Reprinted figure with permission from [124]. Copyright (2002) by the American Physical Society.)

4.2.5 Surface coating methods for microchannel walls Similar to the conventional capillary electrophoresis, the surface properties of the internal walls of a microchip have a significant impact on the performance of the separation. Surface coating is usually required to reduce the electroosmotic flow (EOF) and to minimize the analyte-wall interactions, both of which deteriorate the separation efficiency by introducing band-broadening during gel electrophoresis. The nonuniform surface charge resulting from the material itself or/and the micromachining process, as well as the nonspecifically absorbed DNA molecules over the runs, affects the EOF during electrophoresis, and thus severely deteriorates the reproducibility. In addition, the large surface-to-volume ratio in a microchip makes it even more important to suppress the interaction between DNA and the internal wall, since the small volume of samples can easily be depleted, and hence detection sensitivity impaired. Surface coating strategies can be divided into two categories: dynamic coating (or physical adsorption) and permanent coating (or chemical modi-


fication) [125-129]. Compared to the often time-consuming and laborintensive chemical modification, dynamic coating is a simple, fast and low cost option, especially for disposable plastic microchips. But the permanent surface coating is usually more uniform and stable. Following the same categorization as microchip fabrications, both surface-coating chemistries will be reviewed in the following section based on different substrate materials. 4.2.5.1 Glass/silicon chip surface coatings

Both the dynamic and permanent (or physical and chemical) coating methods that have been well developed for fused-silica capillaries during the “capillary era” are fully transferable to glass/silicon microchips simply due to the fact that they are the same material and share the same surface chemistry. Various polymers, such as PDMA, PEO, PVP, PVA and HEC, have been identified not only as a sieving matrix, but also as a dynamic coating materials in both DNA fragment sizing and long-read DNA sequencing applications [125]. The “self-coating” is simply realized by physical adsorption and coating the microchip can be easily carried out by pumping the polymer matrices through the microchannel. Compared to the multi-step chemical modification procedures, dynamic coating is more amenable for automation and electrophoresis integration. On the other hand, the most common surface coating for glass microchips is still chemical modification because of its robustness and well-documented history. It involves the use of the classic Hiertén protocol [130] or its modified version, in which a neutral polymer layer is covalently bond to the glass/silicon surface activated by a standard silanization chemistry. For example, the protocol of covalently bonding the polyacrylamide through the γmethacryloxypropyltrimethoxysilane was optimized on the surfaces of glass microchips for both fragment analysis and sequencing applications [131, 132]. Chemically modifying the glass surfaces with a thin-layer of PVA coating was also demonstrated [133]. 4.2.5.2 Polymer/plastic chip surface coatings

PDMS is one of the most popular substrate materials for microfluidic chips. Unfortunately, the native PDMS is not ideal for electrophoresis due to its inherent strong surface hydrophobicity, because of which the microchannel is very difficult to wet and air bubbles are easily generated. Additionally, the strong tendency for analyte absorption produces unstable

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EOF. Numerous dynamic coating and chemical modification strategies have been proposed to tailor the surface properties of PDMS for different applications [127]. HPC and HPMC were used as the PDMS self-coating sieving matrices for dsDNA fragment analysis in a few early reports, though the authors did not clearly indicate the dynamic coating properties of the HPC and HPMC [134, 135]. PDMS surfaces can be chemically modified by silanization or UV grafting. When using silanization, PDMS surfaces are treated with oxygen plasma or Tesla coil to oxidize the Si-CH3 group to Si-OH group, followed by the similar silanization chemistry applied on glass/silicon surfaces [136, 137]. When using UV grafting, PDMS microchannels are filled with different monomers, including acrylic acid, acrylamide and PEG-monomethoxyl acrylate, etc. After exposure under a mercury lamp, these monomers are grafted to the PDMS surface to yield hydrophilic surfaces and produce stable EOF [138-140]. Surface modification on thermoplastics, including PMMA, PC, and COC has been substantially studied to make them ideal substitutes to glass microchips, offering not only the low cost, but also the high performance for electrophoresis [125]. However, each thermoplastic material has its unique functional groups at the surface. Unlike the universal silanization chemistry applied to glass and PDMS surfaces, a different chemistry has to be designed and optimized for each surface and those polymers may not be compatible with some of the organic solvents used during chemical modification. Under such circumstances, dynamic coating is especially attractive simply because the usually tedious chemical modification does not harmonize with the low cost and disposable nature of those chips. Several dynamic coating methods have been investigated for the surfaces of the thermoplastics. Hydrophilic neutral polymers including PEG, HEC, HPMC, and methylcellulose (MC) have been tested for PMMA surface dynamic coating. For example, HPMC and Pluronic polyol F127 (PEO99PPO69PEO99) were used as self-coating sieving matrices for dsDNA separation in PMMA microchip by Xu et al. [141] and Song et al. [94], respectively. DNA sequencing and STR genotyping were successfully demonstrated in Zeonor (poly(cycloolefin)) microchip dynamically coated with 2% poly(dimethylacrylamide/diethylacrylamide) [33].


Although not aligned with the low cost nature of the disposable plastic microchips, to demonstrate the feasibility of using those thermoplastic materials for electrophoresis, many chemically modification strategies have been pursued [142, 143]. The chemical modifications allow some improvement of the analytical performance of the plastic microchips but they are still not good enough for the high-resolution applications like DNA sequencing.

4.3 Parallelization in Microchip Electrophoresis One of the easy ways of implementing multiplexing on an electrophoresis microchip is to facilitate channel parallelization, namely “microchip array electrophoresis” chips. Single-channel electrophoresis microchips have already shown clear advantages over the conventional slab gel or capillary electrophoresis with respect to analysis time, separation efficiency, and sample consumption, etc. However, microchips’ superiority will not be fully revealed without high sample throughput being realized on them. Fortunately, one of the most attractive features of the microfabricated devices is that intricate, ultra-dense channel arrays can be easily fabricated onto a single chip without significant increase in time and cost as compared to the single channel microchip fabrication. In 1997, Woolley et al. first demonstrated the feasibility of genotyping 12 samples simultaneously on a glass microchip with a rectilinear network of 12 channels [144]. Simpson et al. then expanded it to 48 channels with each channel capable of analyzing two samples sequentially. The system allowed an analysis of 96 samples in 8 minutes [145]. To further increase the density of the channel and address the limitation of a roster optical scanner, Shi et al. developed a rotary confocal fluorescence scanner together with a microchip that consists of 96 channels radially distributed on a 100mm-diameter wafer [146], which have been used to demonstrate a series of clinical genetic testing. The radial design was further advanced with the presentation of a 384-channel array on a 200mm-diameter wafer [147]. With 80 mm effective separation length, this microchip demonstrated genotyping of 384 individuals in only 325 s for the common hemochromatosis-linked H63D mutation in the human HFE gene, corresponding to a throughput of more than 1 sample per second. By multiplexing also in the detection aspect, for example spectral multiplexing, this format was expected to yield a throughput of 147,000 samples/day, a 20-fold increase over the commercial 96-capillary systems.

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While microchip array electrophoresis gained momentous successes in genotyping applications, several research teams, including Mathies’s group at UC Berkeley and Whitehead BioMEMS laboratory at MIT led by Ehrlich, were engaged in developing microchip based high throughput sequencing instruments in response to the call for “ultra-low-cost sequencing techniques” or “personal genome project”. The first DNA sequencing in a microchip array was demonstrated by Liu et al. in 2000 [148]. The 16 simple-cross channels were fanned out on a 100mm-diameter wafer. The system routinely yielded more than 450 bases in 15 min in each of the 16 channels. At almost the same time, Simpson et al. and Backhouse et al. both fabricated 48 straight channels on glass substrates [149, 150] for DNA sequencing. These micro channel array designs are a complete miniaturized version of a conventional capillary array system, on which electrokinetic sample injection along the separation channel rather than the cross-T injector was used. The typical advantages associated with the microchip, such as the shorter separation length and the faster analysis, were not fully realized in these works. Based on the theoretical prediction and experimental investigation on the turn geometries for minimizing band broadening in microchip electrophoresis [54, 151, 152], Paegel et al. introduced a “pinched turn” to extend the separation length from 55 mm to 159 mm on a 150mm-diameter wafer, which is necessary in DNA sequencing for single base resolution. An average read length of 430 bases at 99% base-call accuracy was obtained in 24 min, equivalent to a sequencing rate of 30 bases per second [153]. By replacing the LPA sieving matrix with Nanogel, a read length of about 500 bp at 98% calling accuracy was obtained in 25 minutes [92]. Researchers at Whitehead BioMEMS lab did a systematic study on the important parameters that affect the DNA sequencing performance in microchips and fabricated 32 identical 40-cm long microchannels on a 25 cm x 50 cm glass plate. This device achieved a read length of 800 bases, but at the cost of a longer separation time (80 min) [154-155]. While the race of maximizing the array density continues, the final trophy went to the prototype of BioMEMS-768 high throughput sequencing system [31, 156]. Two 384-channel plates were cycled alternatively between electrophoresis and regeneration. In each run, a total of greater than 172,000 bases, 99% calling accuracy was generated in ~70 minutes, corresponding to a total of a 4 megabase throughtput per day. For comparison, it takes about an hour for the latest ABI 3730xl 96-capillary DNA Sequencer to generate 67200 bases (700 bases each capillary) at an accuracy of 99% in a standard opera-


tion condition, corresponding to ~1.6 megabase per day. With the significant reduction in reagent consumption, microchip electrophoresis becomes one of the most promising technologies to deliver reasonably low cost whole genome sequencing (~$100,000 genome). A summary of multiplexed electrophoresis microchips is given in Table 4.1. Table 4.1 Summary of multichannel electrophoresis microchips for DNA sequencing Refer- # of Pattern ence Channels

Separa- Separa- Sample tion tion /Template Length Matrix (mm)

Accu- Read Time Length (min) racy (%) (bp)

[148]

16

Double T 70-76

LPA

Unknown

99

450

15

[150]

48

Straight

456

POP-6

BigDye Sample

98

640

150

[149]

48

Straight

100

LPA

M13mp18 97

400

[155]

32

Cross

400

LPA

M13mp18 98

800

78

[153]

96

Double T 159

LPA

M13mp18 99

430

24

[92]

96

Double T 159

NanoM13mp18 98.5 gel

500

24

[31]

768

Cross

M13mp18 98.5

580

~70

370450

LPA

4.4 Integration in Microchip Electrophoresis for Genetic Analysis 4.4.1 Sample preparation on microchip Sample preparation steps prior to electrophoresis, which include DNA extraction, amplification, and purification etc., are extremely critical for high

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quality genotyping or sequencing data. The conventional sample preparation procedures generally involve a series of benchtop instruments doing off-chip processing and the sample volumes involved are usually several orders of magnitude larger than what is needed for the application on a microchip. Those processing procedures are usually cumbersome and often limit the quoted advantages of microchip electrophoresis, such as low reagents consumption, low cost and high throughput. Novel methods for onchip sample preparation with the intention of integration with the downstream microchip application have been proposed and demonstrated. Sequence specific DNA fragment enrichment is generally required in genetic analysis. Since its inception, PCR (polymerase chain reaction) is the most powerful tool to amplify the target DNA fragment millions to billions of times. Implementing PCR on a microdevice has been of great interests to scientists and engineers. Microchip PCR devices can be considered having two categories: stationary-chamber PCR and continuous flow PCR. The stationary chamber PCR device is basically a miniaturized thermocycler. Nanoliter scale PCR reactions have been demonstrated in this format [157]. The continuous flow format, which limits the thermal mass to the sample solution instead of the whole devices, generally provides faster PCR [158, 159]. A few recent reviews have summarized the progresses of microchip PCR [160-163]. This chapter will only mention the PCR microdevices that are or were intended to be coupled to microchip electrophoresis. Mathies group developed a monolithic, nanoliter DNA amplification and electrophoresis system with a microfabricated thin film heater and resistive temperature detector (RTD) [164-166]. Landers group demonstrated a fast non-contact PCR reactor by employing an IR temperature control system [167, 168]. In DNA sequencing, amplification of various lengths of DNA fragments is typically carried out by a Sanger cycle sequencing reaction, a similar process as PCR with the difference being the random termination of the chain extension when ddNTP incorporated. Therefore, any microchip design that is suitable for PCR can be applied to Sanger sequencing reaction as well. Both stationary and continuous flow formats have been implemented for Sanger cycle sequencing reactions. The Mathies group used the abovementioned configuration to complete the Sanger extension on a microchip [60]. Soper et al. demonstrated Sanger sequencing with a 62 nL reactor coupled directly to a capillary gel column for DNA separation [169]. Compared to PCR, Sanger extension is much slower because the polymerase is altered to improve the incorporation efficiency of the phosphate analogs, e.g. ddNTPs or dye labeled ddNTPs. By improving the heat transfer with a


continuous flow microchip, Wang et al. pushed the Sanger reaction speed to the enzymatic limit [170]. A purification step is generally required before the PCR amplification of DNA from a crude biological sample or after the Sanger cycle sequencing reactions in a sequencing experiment. The purification before PCR amplification is to remove the inhibitors and/or pre-concentrate the DNA templates to ensure the efficiency of PCR. The purification of Sanger extension reaction products is routinely carried out before loading the sample onto a capillary or a microchip to remove the excess amount of DNA template, unincorporated nucleotides, and salts. Those impurities with high electrophoretic mobility could adversely affect the sequencing results including the read-length and the separation resolution. The conventional DNA purification methods are ethanol precipitation or solid phase extraction. Those are time-consuming and involve multiple steps that are not feasible to be incorporated on a microchip. Alternative approaches have been investigated for better purification quality and more importantly for an easy integration into a micro-total-analysis system. The Landers group explored solid-phase extraction (SPE), which relies on DNA adsorption to silica resin in the presence of a chaotropic agent, to extract, purify and pre-concentrate DNA from raw samples from human, bacteria, and virus etc. [171-173]. Xu et al. purified the Sanger extension fragments in a UV modified polycarbonate microchip with an adapted solid-phase reversible immobilization (SPRI) chemistry [174], which was first demonstrated with carboxylated magnetic microbeads by Hawkins et al [175, 176]. Paegel et al. demonstrated a low viscosity hydrogel matrix for selectively hybridizing and releasing the Sanger extension products to achieve the sample pre-concentration and purification [177]. 4.4.2 System integration Developing a fully integrated DNA analysis system on a single chip has been the ultimate goal of many researchers since the inception of the concept: lab-on-a-chip [178]. Typical sample transfer methods using conventional pipette tips or automatic sample dispensers at microliter scale obviously degrade most of the advantages associated with the microchip. The true power of the miniaturization will not be fully revealed without system integration. Although the microchip electrophoresis device alone has shown superior separation performance and the related sample preparation units have also shown great promise in processing nanoliter volumes of

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sample, integrating various functions together and streamlining the total operation on microchips still represents great challenges to researchers across all disciplines. Thus far, only a handful of integrated microchip genetic analyses have been demonstrated. The first integration, demonstrated by Burns et al. in 1998, reported an integrated nanoliter DNA analysis device that was capable of performing sample injection, amplification or digestion, and electrophoretic separation [179]. The liquid manipulation units, thermal control units, in-situ crosslinked polyacrylamide gel electrophoresis units, and photodiode detectors were all fabricated on a silicon substrate by conventional micromachining techniques (Fig. 4.5). One of the keys to the success of the proof-ofconcept device was the hydrophobic patch valves and capillary force pumps for nanoliter scale liquid manipulation. However, it is challenging for surface tension based pumps and valves to manipulate different fluids (e.g. real biological samples, different buffers) in more complex geometries.

Fig. 4.5 (Top) Schematic of integrated device with two liquid samples and electrophoresis gel present. (Bottom) Optical micrograph of the device from above. Wire bonds to the printed circuit board can be seen along the top edge of the device. The pressure manifold and buffer wells that fit over the entry holes at each end of the device are not shown. (From [179]. Reprinted with permission from AAAS.)


Following the first integrated “Lab-on-a-chip” device, there have been a few other attempts of integrating the sample preparation units with the electrophoresis units. The progress was partially hindered by the lack of interconnecting components, e.g. micropumps, micro-valves, and micromixers, for seamlessly interfacing various functional units and assembling them into a fully functionized and miniaturized system. By utilizing the pneumatically activated valves and pumps, two research protoytpes of miniaturized and fully integrated electrophoretic systems targeting real biological samples were finally revealed in 2006, nearly one decade after the first proof-of-concept lab-on-a-chip device. Mathies et al. incorporated all three Sanger sequencing steps, thermal cycling, purification, and electrophoretic separation, into a fully integrated and automated format (Fig. 4.6) [60]. With a 30-cm long separation channel filled by linear polyacrylamide gel, a read length of 556 bases with 99% accuracy from 1 fmole DNA template was successfully demonstrated. The reagent consumption and the cost of the analysis, was reduced by ~100 fold (from 20 µL to 250 nL), a dramatic improvement simply because of the removal of the off-chip sample transfer steps. The multi-layer glass-PDMS hybrid lab-on-a-chip assembly involved using several generic components that could perform the essential individual process, so it could be potentially used in a wide range of bioanalytical and biomedical applications. Compared to the pure DNA template used by Mathies et al., Landers et al. demonstrated a “sample-in-answer-out” device which was capable of accepting crude biological samples, such as blood and other bodily fluids [180]. Two sample preparation units, including a solid phase DNA extraction (SPE) unit to carry out on-chip DNA purification and a PCR amplification unit, were integrated to an electrophoretic analysis unit through a combination of several recent innovations in microfluidics, such as differential flow resistances, elastomeric valves, and laminar flows. The integrated microsystem could screen pathogen infection from a very small (sub-microliter) volume of crude samples in less than 30 minutes, as demonstrated by the identification of the presence of Bacillus anthracis (anthrax) in 0.75 µL of whole blood from infected but asymptomatic mice, and of Bordetella pertussis in 1 µL of nasal aspirate from a suspected human patient.

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Fig. 4.6 Integrated DNA sequencing bioprocessor. (Reprinted with permission from [60]. Copyright (2006) National Academy of Sciences, U.S.A.)

It has been a long way from the birth of the concept of “lab-on-a-chip” to the first published successful research prototypes. It may take another few years before the turnkey “sample-to-answer” micro-total analysis system eventually comes to the real world to be used for point-of-care clinical diagnostics or at the crime scene for forensic analysis. With the continuing growth of microfluidic technology and little expectation for such a technology to reach a plateau, scientists and engineers are taking on the daunting challenges and getting closer and closer to reaching their ultimate goal.

4.5 Commercial Microfluidic Instruments for Genetic Analyses An ultimate validation of a scientific idea may be its realization into a commercial product that subsequently benefits society. Since the conceptual birth of “lab-on-a-chip” almost two decades ago, there has been tre-


mendous interest and incredible technological advancement over the years, as evidenced by the enormous amount of publications partially summarized in the previous sections. Following its success at the research level and considering the great promise it holds for almost every life science and health care area, microfluidics technology is also expected to have widespread commercial use. The main drivers that promote microfluidics products into the commercial markets are the inherent merits of miniaturization mentioned previously: rapid analysis time, reduced sample volume and reagents cost, and great labor savings offered by system automation and integration. The reduction of assay time from hours to minutes or even seconds by using the microfluidics technology is the booster of the emerging microfluidics market. The micro, nano, or pico liter of volume needed for analysis and the same scale of waste produced on a microchip system is another market driving force. The cost per assay is dramatically reduced by ten or more fold due to the lower reagent consumption. In addition, its flexibility of interfacing with other methods and technologies for automation allows a total analysis to be carried out without user intervention, resulting in a great labor savings. The goal of this section is to summarize the major commercial microfluidic products on the market for genetic analysis and to give a general overview of the rising microfluidic markets. The authors have found that the microfluidics products evolve very rapidly with the changing market. Therefore, rather than concentrating on specific products, we will guide the readers through different companies by their unique technologies and then introduce their products based on such technologies. 4.5.1 Commercial microchip electrophoresis instruments for genetic analysis A summary of commercial microfluidic instruments that specialize in microchip electrophoresis will be first given in this section followed by a few other microfluidic products that carry out a broader scope of genetic analyses including sample preparation and micro-array applications. A few examples of companies who are making microchip electrophoresis products are: Agilent Technologies, Bio-Rad Laboratories, and Caliper Life Sciences. Their current products are 2100 Bioanalyzer, Experion Automated Electrophoresis System, and Labchip 90, respectively. All

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three products are using the same LapChip® technology developed by Caliper Life Sciences. The LapChip® technology was described by Bousse etc. [181]. Briefly, it automates steps involved in gel-based electrophoresis for separation of biomolecules including DNA, RNA, and protein. Electrophoretic separation, analyte staining and detection are integrated on a single chip and the process is automatically repeated for multiple-sample analysis. DNA sizing and quantification of ten samples can be completed in 30 min on a chip, a dramatic improvement over conventional capillary instruments. Agilent 2100 Bioanalyzer was the first (in 1999) commercialized microfluidic product based on Caliper’s LapChip® technology through collaboration between the two companies [182, 183]. Similarly, Bio-Rad launched the Experion Automated Electrophoresis System in 2004 using the same technology by collaborating with Caliper as well [183]. Bioanalyzer and Experion are considered as low-throughput miniaturized electrophoresis systems. The number of samples that can be analyzed depend on the number of wells on a chip and both systems currently analyze ten DNA samples per chip. Caliper’s own microchip electrophoresis product, LabChip90, is a high-throughput alternative to the Bioanalyzer and Experion systems. Different from the other two automated electrophoresis systems, LabChip90 can be coupled with 96 or 384 micro-titer plates, automatically draw samples from a well-plate sequentially using Caliper’s proprietary “sipper” sampling system, then analyze each sample under a typical gelelectrophoresis condition (www.caliperls.com). The system automates electrophoresis, detection, and data analysis for processing thousands of samples per day. A variety of chips and reagents designed for different applications are offered along with the instrument as consumables. The applications for DNA analysis include genotyping, micro-array production, and gene expression studies. The commonality of the above mentioned LapChip® technology-based microfluidic instruments are that they all integrate and automate certain functionality such as separation, staining, destaining, and detection in the workflow. However, the cumbersome sample preparation steps involved in a complete DNA analysis (e.g. from crude samples) are all done manually prior to sample loading onto the instruments. Those products are similar to the conventional bench top capillary or slab gel systems as to the application, but they provide faster analysis speed and higher throughput. These products have been well received by the customers and the success of these microfluidic products has helped the companies gaining big market shares in the microfluidic industry. The revolutionary separation speed


and data quality of the first microfluidic instruments on the market are evident enough for researchers to realize the power of the new generation of separation platform. 4.5.2 Integrated microfluidic instruments for genetic analyses The possibility of the many steps of conventional assays being streamlined into a single process and in the nanoliter scale is very attractive. An ideal lab-on-a-chip system should automate and integrate all steps involved in an analysis on one platform. The “sample-in” and “result-out” format of Lab-on-a-chip or Micro-total-analysis system reduces analysis time, risk of sample loss or contamination, and is a perfect replacement for bulky, expensive laboratory robots. In this section, we introduce a few companies who are developing integrated genetic analysis instruments using their proprietary microfluidics technologies. Cepheid is one of the pioneers in commercializing the integrated genetic analysis system (www.cepheid.com). By using its smart cycler technology (I-Core®) that integrates thermal cycling with micro-optics and electronics, Cepheid offered customers two versions of PCR-based genetic analysis systems: the SmartCycler® and the GeneXpert®. These products have been used in rapid genetic screening for infectious diseases, cancer, and bio-terrorism detection. Cepheid offers series of testing and reagent kits to go with the instruments to amplify and identify a specific DNA sequence associated with a disease or bio-threat agents. SmartCycler® is a real-time PCR system that can provide answers from a processed biological sample in 30-40 minutes. The GeneXpert® system integrates sample preparation with PCR amplification and real time detection for a fully automated genetic analysis. It purifies, concentrates, amplifies and detects targeted nucleic acid sequences in less than 30 minutes from a raw sample. It uses a disposable microfluidic cartridge together with a special designed PCR reaction tube. After raw specimens lysed by ultrasonic force, the released DNA and reaction mixtures are pumped through chambers inside the cartridge by a syringe drive and a rotary drive and finally reach the reaction tube for thermal cycling and detection. The throughput of the system can be scalable up to 16 modules at this moment with each module carrying out one test (www.cepheid.com). Microchip Biotechnologies (MBI) is another microfluidic company that is developing a “sample-to-answer” format of genetic analysis system (http://www.microchipbiotech.com). Specifically, they are developing an

148 Zhu and Wang

integrated microfluidic system that includes sample preparation and analysis for DNA sequencing. The two key technologies: Microbead Capture technology and Microscale on-chip valves (MOV™) technology focus on the interfacing between the macro- and micro- world. The Microbead Capture technology uses coated magnetic beads to capture and concentrate specific biological molecules from a large amount of complexed sample matrices into micro- or nanoliter size. It serves to bridge large volume sample with the downstream micro-volume analysis. MOV™ technology is one of the on-chip “valving and pumping” designs that enable the successful micro or nanofluidic mixing and fluid transferring within the intricate microfluidic network. The on-chip valves and pumps are the key components that allow system miniaturization, automation, and integration. Combining these two key technologies and together with a robotic workstation, MBI has developed an automated sequencing sample preparation instrument, Apollo 100. This instrument automatically sets up on-chip Sanger cycle sequencing reaction and then perform sample clean up for the downstream detection. During the automated process, robots move the DNA template and the reaction mix onto the microchip, MOV™ assist with pumping and mixing of samples for thermocycling and finally the magnetic beads come into place to cleanup the sequencing reaction products. Not connected with a DNA sequencer and also using pre-processed DNA template, the system is not fully automated in terms of sequencing from a raw sample. However, the Apollo 100 will be beneficial to those who already own capillary DNA sequencers but perform cumbersome sample preparation steps manually (http://www.microchipbiotech.com).. Fluidigm introduced the technology of “integrated fluidic circuits (IFCs)” to the microfluidic industry and launched the BioMark™ product line based on such a technology for variety of genetic analysis (www.fluidigm.com). IFCs are microdevices fabricated with networks of fluid-control valves (NanoFlex™ valve) and interconnected channels, in which the movement of the biological sample and reagents can be regulated. The core of this technology is the NanoFlex™ valve which consists of layers of elastomers that deflect under pressure to form a seal. Using NanoFlex™ valves’ compact size and precise fluidic control, Fluidigm established high throughput genetic assay platforms, the Biomark™ 48.48 Dynamic Array and Biomark™ digital arrays. The Biomark™ products offer real-time PCR and genotyping at a high throughput together with accurate DNA quantification (www.fluidigm.com). If microscale liquid handling and micro-array printing are also considered within the realm of the microfluidics, Akonni Biosysem may need to be


mentioned for its TruArray® technology (www.akonni.com). It is an integrated microarray technology that screens a biological sample against hundreds of diseases markers simultaneously and gives an answer in less than 30 min. An array of the micro-gel-drops is precisely loaded onto a plastic microfluidic slide, size of a credit card. Each micro-gel-drop carries out an individual PCR reaction, tailored for a specific diagnostic test. Coupling this on-chip PCR with precise microfluidic control, the portable TruArray® system enables hundreds to thousands of diagnostic tests performed on a clinical sample in a short period of time (www.akonni.com). There are also dozens of other companies that hold patented or licensed microfluidics technologies with their products spanning over a broad spectrum of applications. A table (Table 4.2) that summarizes the current active microfluidic companies and their major technologies for genetic analysis is given below including some examples that have not been discussed in the text. The authors do not intended to provide a complete list of all the microfluidics companies and all their microfluidic products. Even at the time of this book being published, dozens of new companies are getting started and new product ideas being introduced. Table 4.2 Some microfluidic companies and their commercialized microfluidic instruments for genetic analysis Company

Akonni Biosystems www.akonni.com Agilent Technologies www.agilent.com Bio-Rad Laboratories www.bio-rad.com

Technology TruArray™ with microfluidic liquid handling) LabChip® for gel electrophoresis LabChip® for gel electrophoresis

Genetic tion

Applica-

Clinical molecular diagnosis Electrophoresis and quantification Electrophoresis and quantification n

Bio-Trove www.biotrove.com

OpenArray™

Genotyping, time PCR

Caliper Life Sciences www.caliperls.com

LabChip® for gel electrophoresis

Cepheid www.cepheid.com

Smart Cycler technology (I-Core®)

Electrophoresis and quantification, Sequencing Real-time PCR, Clinical diagnosis

real-

Product for genetic analysis TruArray™ 2100 Bioanalyzer Experion Automated Electrophoresis System OpenAray™ SNP Genotyping System LabChip® 90

GeneXpert SmartCycler

150 Zhu and Wang Table 4.2 cont. Fluidigm www.fluidigm.com

Integrated Fluidic circuits (IFCs) NanoFlex™ valve\

Handylab www.handylab.com

Microthermal Circuit

Microchip Biotechnologies www.microchipbiotec h.com Nanogen www.nanogen.com

Microbead Capture Technology, Microscale on-chip valves (MOV™) Nanochip® Electronic Microarray

Genotyping, gene expression profiling, quantification, diagnostic Real-time PCR, Clinical diagnosis DNA Sequencing

SNP analysis, STR analysis, mutation analysis

BioMark™ Dynamic array BioMark™ Digital array Jajuar®, Lynx® (under development) Apollo 100

NanoChip® 400 system (product closed in 2007)

4.6 Microfluidic Markets and Future Perspectives The market for microfluidics/lab-on-a-chip products is growing rapidly with the technologies developed in the field. With the significant progress and the high expectation in both the technology and the market, customers have begun to accept the microfluidic products . One of the hurdles the customer has to overcome before the microfluidic products can stand firmly in the market is the replacement of existing well-functioning and highly invested large-scale robotic systems with the maybe even more expensive microfluidic products. In addition, products that come out of new technologies often have risks associated with them, even though many considerations are put into improving the products’ reliability and cost. However, with a considerable amount of investment from both government and private sectors continuing to go into the R&D laboratories of numerous start-up and well-established companies to develop microfluidic instruments, the market potential for microfluidic industry seems to be rising. More and more customers may gradually realize the long-term savings effect brought by the microfluidic technology and decide to invest in it, especially in the cases where reduction of cost and limited amount of sample are prominent. In the meantime, many microfluidics manufacturers are striving for greater accuracy, improved throughput, increased automation, and better robustness. There seems to be quite potential market spaces for this continuing maturing technology. Many are expecting the microfluidic


market to flourish in the next a few years as the novel technology gains more acceptances among the research labs around the world. The continued development of microfluidic industry, on the other hand, will depend upon the successful realization of miniaturization and integration. Performing all steps of a biological assay on a single microchip is very attractive for its obvious advantages in terms of speed, cost, and automation. Currently, a lot of robotic workstations are being used as enabling tools at the interface of microchips and sample processing steps. Although they have provided great productivity and accuracy, such complicated and expensive equipment may not hold up long under the increasing demands for portable and fieldable integrated total DNA analysis systems. For that reason, the microfluidics industry may aim at creating new genetic analysis platforms that incorporate greater functionalities but still with a small footprint. As indicated by the latest research publications, as well as a few products listed above, one of the obvious research trends is to produce a “sample-in, answer out” high-capacity microfluidic product. Microfluidics manufactures are gaining momentum through continuous technological breakthroughs that are able to address the unmet market needs. The gap between where the technology is and where the market needs it to be continues to get smaller and smaller.

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Chapter 5 Microfluidic Systems for Protein Separations

Anup K. Singh Biosystem Research Department, Sandia National Laboratories, Livermore, CA 94551 Correspondence should be addressed to: Anup K. Singh ([email protected])

Keywords: Microfluidic protein separation, microfluidic liquid chromatography, multi-dimensional separation, electrophoresis

Abstract Proteins are the molecules that carry out virtually all functions in living cells. Consequently, numerous methods have been developed to separate and analyze proteins for their identification and quantification. Microfluidic devices have attracted significant attention for proteomic analysis owing to a number of advantages they offer over conventional methods including improved speed of analysis, higher resolution, increased multiplexing, and the ability to analyze minute amounts of sample.

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5.1 Introduction Emerging research areas in biology and biotechnology, such as genomics, proteomics, and structural biology, increasingly require a large number of experiments performed in a smaller amount of time. Moreover, in most instances, these ever-increasing number of experiments need to be performed using a limiting amount of starting biological sample. These needs require a scaling down of the analysis methods. In response to this trend, analogous to the integrated circuit (IC)-chip revolution, “microfluidic chips” are starting to transform the field of biochemical analysis. While the search for “the killer application” of microfluidics continues, microfluidic chips have come a long way over the last decade and, in addition to academic research applications, they are being used in many commercial devices for analysis of DNA, RNA, proteins, and cells. In this chapter, we will focus on application of microfluidic chips for protein separations and proteomics.

5.1.1 Advantages of microfluidic chips for protein separations There are many reasons for ever-increasing applications of microchips for protein analysis as summarized below. Faster separation: Adaptation of protein analysis to a chip leads to a 10 100-fold increase in the speed of analysis. The key reason for this is the scaling down of analysis or separation dimensions compared to conventional counterparts. For example, the separation distance covered in slabgel protein electrophoresis is typically 6-10 cm. In a chip, the length of a separation channel could be less than 0.5 cm and hence, the same sample can be analyzed 12-20 times faster. Of course, this also requires a scaling down of sample size, but in many applications this is an additional advantage. For electrokinetic separations, there is another factor that improves the speed - the ability to apply higher electric fields with everything else remaining constant. In slab-gel electrophoresis, Joule heating limits the applied electric field to less than 250V/cm. In a glass chip, since glass is a good heat conductor and microchannels boast high surface area-to-volume ratios, fields as high as 750V/cm are routine, leading to another 3-5 times improvement in speed. With the chip format, improvement in speed enables analysis of a larger number of samples per unit time, as well as quicker sample-to-answer times compared to conventional methods.

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Smaller sample size and reagent volume: Another key advantage offered by microfluidic systems is that they require, and are capable of handling, a very small amount of sample and reagent for each analysis - a few tens or hundreds of nanoliters. These volumes are impossible to analyze in conventional microtiter plates or vials as they will evaporate within seconds. This advantage also results from the scaled-down dimensions offered in a microchip with sealed microchannels. In addition, the small size of the chips themselves, make it possible to reduce the footprint of the analysis device. It is anticipated that in the not too distant future, we will have a personal or portable protein analyzer based on a protein analysis chip.

5.1.2 Limitations of microfluidic chips in proteomics applications The key limitations of microchips are side-effects of the advantages they offer. For example, while miniaturization enables faster analysis, it reduces the peak-capacity of chip-based separations. Current microchips cannot perform the equivalent of a conventional 2-dimensional separation on a complex sample such as cell lysate or blood serum. For similar reasons, chips also cannot handle complex dirty samples. Limited peak capacity makes it hard to resolve peaks if there are too many components present or if one component is present in huge excess. Another factor, resulting from the small cross-sections involved, is that it is relatively easy to clog the channels with particulates or aggregates in the sample. It is also very easy to foul the separation channels, which reduces run-to-run reproducibility and limits reuse of the chip. Scaling down of dimensions results in a smaller path-length for optical detection of analytes as well. So while limit of detection (LOD) in terms of amounts (or moles) is superb in chips, the concentration sensitivity can be very poor. This makes detection methods such as absorbance unsuitable for microfluidic analysis. One typically has to use fluorescence, chemiluminescence or electrochemical detection methods with chips.

5.1.3 Substrates used for proteomic analysis Glass, fused-silica, or quartz have been the most widely used substrates and have been commercialized by companies such as Agilent, Caliper and Bio-Rad. Other vendors for these chips are Micronit and Micralyne. Plastic

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chips have also been developed - owing to their inexpensiveness and simple fabrication. The key criterion used for suitable polymers to fabricate a chip is the requirement that they are transparent in spectral regions of interest for available detectors. Hence, the possible choices are acrylates such as Polymethyl methacrylate (PMMA), Polydimethylsiloxane (PDMS), cyclic olefin copolymer (COC), and polycarbonate (PC). The most widely-used polymeric substrates are PMMA and PDMS. In the early days, researchers had to find fabrication facilities in-house or through a collaborator who had access to one. This really limited the places where microfluidic research could be done. Now, with wider acceptance of chip-based analysis, there are a number of commercial vendors that provide standard as well as custom chips for virtually any application.

5.2. Microfluidic Chips for Protein Separation Most of the early applications of chips for proteomic applications utilized open-channel separations. The first successful adaptation of a protein analysis technique to the microchip was capillary zone electrophoresis (CZE) because it is perhaps the simplest and easiest method to adapt to a chip. Other protein separation applications adapted to microfluidic chips include: capillary gel electrophoresis (CGE), Sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE), isoelectric focusing (IEF), micellar electrokinetic chromatography (MEKC), isotachophoresis (ITP) and open-tubular chromatography. Figure 5.1 lists the protein separation techniques that have been adapted to chips.


Capillary Electrophoresis Gel-Based: CGE PAGE & SDS-PAGE Open-channel: CZE ITP IEF MEKC

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Liquid Chromatography Ion Exchange Size Exclusion Reversed-Phase Electrochromatography Hydrophobic Interaction Affinity

Multi-Dimensional Techniques LC-MS CE-MS LC-CE 2D Gel Electrophoresis

Fig. 5.1 Protein separation techniques adapted to microfluidic chips.

5.2.1 Microchip-based electrophoretic techniques 5.2.1.1.Capillary zone electrophoresis of proteins

Capillary zone electrophoresis (CZE) separates analytes based on their charge/mass ratio and is carried out in a capillary filled with an aqueous buffer. CZE offers extremely high resolution and hence, has been widely used for the separation of proteins and peptides. CZE has been adapted to a planar microchannel format by a number of researchers [1-10]. Chip-based CZE allows better control of sample introduction and leads to better performance in terms of speed and efficiency over the conventional capillarybased separation. Microchip-based CZE has also been multiplexed and integrated with other forms of separation to enable multidimensional separations. Chips for CZE of proteins, similar to capillary-based separations, require that channel walls be coated covalently or dynamically to reduce non-specific adsorption of analytes. Proteins can adsorb non-specifically on glass surfaces due to electrostatic forces (for cationic proteins or even anionic or neutral ones with cationic domains) or hydrophobic interactions. Fortunately, many different covalent chemistries or dynamic coating reagents, e.g., polyethylene glycol (PEG) based coating, have been developed for fused-silica capillaries that can readily be used for glass, fused-

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silica or quartz chips [11]. However, for plastic chips, either hydrophilic substrates need to be used or special coatings developed to minimize adsorption [12]. 5.2.1.2. Sizing of proteins

SDS-PAGE is one of the most commonly used techniques for analysis of proteins. It separates proteins based on their size and is typically performed in slab-gels. It is used to determine protein purity, molecular weight and also to identify proteins after a processing step such as affinity separation. The current slab-gel devices require manual loading of samples using pipettes and a typical run is completed in 45 min to an hour. After SDS-PAGE, the gel is stained with a dye, destained to remove excess dye and then imaged in a scanner. These steps take an additional few hours to complete. SDS-PAGE has not substantially changed in the 30 years since its introduction. Adaptation of protein sizing to microchips has overcome many of the limitations of slab-gel electrophoresis by making the separations faster. The most popular chip-based sizing technique is gel electrophoresis using a liquid sieving gel where the gel is made up of linear polyacrylamide or another hydrophilic polymer such as polyethylene glycol [13 - 14]. Unlike slab-gel and CGE in capillaries, the entire process of protein loading, SDSPAGE and detection is integrated and automated. Furthermore, the entire analysis could be completed in less than a minute per sample. This is also an application that as been commercialized as perhaps the most promising microfluidic-chip based device. Three companies, Caliper, Agilent and Bio-Rad, manufacture instruments relying on Caliper’s protein and DNA sizing chips. Slab gel based SDS-PAGE has also been adapted to the microchip format. Han and Singh [15] describe a photopatterning technique to cast in situ crosslinked polyacrylamide gel in a microchannel to perform SDS-PAGE. A fluorescent protein marker sample was separated in less than 30 sec in less than 2 mm of channel length. UV-patterned polyacrylamide gel provides higher sieving power and sample stacking effect, therefore yielding faster separation in a chip. The use of solid polyacrylamide gel instead of liquid sieving matrix in SDS-PAGE also enables easier integration with another separation module.


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5.2.1.3. IEF of proteins

Over the past 30 years, isoelectric focusing (IEF) has grown to be a prominent method of separation and detection in proteome research. The technique separates and focuses analytical species along an established pH gradient based on their different isoelectric points (pI), the pH at which an analyte’s net charge is zero. IEF is commonly used in multi-dimensional separations (i.e. IEF-PAGE) as it provides an orthogonal separation mechanism to size-based fractionation. The earliest microscale adaptation of IEF was done by the Pawliszyn group using a very simple chip containing one channel with an inlet and outlet [16]. The sample was mixed with ampholytes - soluble amphoteric compounds that establish a pH gradient upon application of an electric field - and then loaded into the chip. Voltage was applied to carry out IEF and the entire chip was imaged for detection of the focused bands. Since that initial demonstration, many other articles have also appeared on microscale IEF [15,17-20]. Recently, IEF was achieved on a microchip containing an immobilized pH gradient (IPG), enabling improved pH gradient stability over conventional free-flow methods [21]. The authors established the pH gradient via diffusion of pKspecific acrylamido buffers along a flow-restricted channel. Precise control over boundary conditions and the resulting gradient is achieved by continuous flow of stock solutions through side channels flanking the gradient segment (Figure 5.2). Once the desired gradient is established, it is immobilized via photopolymerization. Rapid (< 20 minutes) isoelectric focusing of several fluorescent pI markers and proteins is demonstrated across pH 3.8 – 7.0 in µIPG’s without the addition of carrier ampholytes, using both denaturing and non-denaturing conditions.

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(a) Low pH

High pH Polymerized Membrane

100µm Immobiline Diffusion

6mm

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(iii)

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Tfer

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OVA

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Fig. 5.2 a)µ IPG fabrication schematics. Gravity was used to provide continuous pressure driven flow from filled reservoirs through the side channels of the glass microchip. A thin polyacrylamide membrane prohibits flow through the 6mm separation channel yet allows equilibration of Immobiline species via diffusion. Entire chip is photopolymerized with UV light following equilibration. b) Composite fluorescence IEF images spanning length of 6mm-long µIPG. (i) Fluorescent pI markers. (ii) AF 647-labeled proteins: Bovine serum albumin (BSA), Transferrin (Tfer), Carbonic anhydrase (CA, bovine), Phosphorylase B (PhB, rabbit muscle), Hemoglobin (Hb, bovine). (iii) AF 488-labeled ovalbumin (OVA) and green fluorescent protein (GFP). Reprinted with permission from [21]. Copyright (2008) American Chemical Society.

5.2.2 Microchip chromatography Chromatography, e.g., high performance liquid chromatography (HPLC), because of its outstanding separation power and versatility, is one of the most common analysis methods for proteins and peptides. There are many


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types of chromatography available, as shown in Figure 5.1, enabled by the wide variety of chromatographic media available. While HPLC studies to date have been implemented in a macroscopic format (in columns with typical capacities and internal diameters of 1 mL and 4.6 mm, respectively) miniaturization presents the potential for several advantages. Improvements have been realized by the advent of microbore and capillarybased HPLC columns, with internal diameters of 100 µm to 2 mm, although injection and detection methods have remained largely unchanged. Significant efforts have been directed towards adapting chromatographic techniques to the microchip format to allow reduction of injection size, a critical advantage when samples are expensive, difficult to generate, or when the scientific question requires minimization of the volume (e.g., single-cell analysis). Reduction of column dimensions also reduces the system flow rate, which leads to improved signal-to-noise when chromatographic separations are connected to concentration-sensitive detectors such as electrospray injection mass spectrometry [22]. A major roadblock in adaptation of chromatography to microchips has been the issue of placement of chromatographic media in the channels. Uniform packing of channels with particles is irreproducible, hard to implement and requires fabrication of shelves and side channels. In the early days of adaptation of chromatography to chips, methods were proposed that obviate the need for packing channels by using open-channel separations. The two examples are- micellar electrokinetic chromatography (MEKC) and open-tubular chromatography. In MEKC, SDS micelles are used as a psuedostationary phase for partitioning of analytes based on their hydrophobicity [23]. While MEKC provides reasonable separation resolution, it does not match the flexibility, reproducibility, peak capacity and resolution of packed-bed reversed-phase chromatography. The biggest drawback of MEKC is that it does not allow the equivalent of gradient elution separations which are so widely used in the resolution of complex peptide and protein mixtures. Open tubular columns with the stationary phase supported on the channel walls is another way to circumvent the use of particles and the accompanying packing problems. Although open tubular columns were first described nearly three decades ago, [24] they have never become popular in liquid chromatography. This is probably because channel widths of 2 µm or less are required to deal with the limited rates of mobile-phase mass transfer in liquid chromatography [25]. Columns with this small diameter are also easily plugged, the loading capacity is extremely small, gradient elution with positive displacement pumps is difficult, and the optical path length used for detection is very short.

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Successful adaptation of packed-bed chromatography to chips was enabled by formation of UV-initiated porous polymers in microchips pioneered by [26] and used by several other research groups [27-28]. In situ casting of polymer monoliths in microchannels reproducibly affords uniform packed beds, therefore eliminating the difficulties associated with packing silica beads and the need for retaining frits. The availability of a wide range of monomers enables critical stationary phase properties such as charge and hydrophobicity to be easily tuned to meet the specific demands of separating many types of analytes. Photopolymerization enables the patterning of a chromatographic media in the microchip, analogous to photolithography, using a mask for optimal design of injection, separation and detection manifolds. These monoliths can be cast in situ in less than 10 minutes, are robust and reproducible with respect to separation characteristics. The microchip in Figure 5.3 was used for analysis of bioactive peptides and amino acids and resulted in separations that were fast (6 peptides in 45 sec), efficient (up to 600,000 plates/m) and reproducible (run-to-run variability 100,000 in human proteome); the wide dynamic range of these proteins; and the variability of gene products (splicing variants, N- and C-terminal truncations, co- and post-translational modifications, etc.). No single separation method has the ability to resolve these

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large numbers (>1000) of components and hence, many multi-dimensional techniques have been developed. The predominant approach in conventional scale experiments has been separation by two-dimensional gel electrophoresis (2-D gel), followed by identification of protein spots using sensitive mass spectrometry techniques. An alternative, and progressively more popular, approach to 2-D gel includes a comprehensive chromatographic separation of the proteolytic fragments derived from intact proteins, followed by mass spectral identification and data base searching. This can be achieved using a two-dimensional liquid chromatography approach whereby peptides are fractionated on a strong cation exchange column, followed by an extended gradient elution on a C18 reverse phase column. Microfluidic systems hold a great promise for realizing multidimensional separations in a single integrated system. To this end, several groups have explored the application of microfluidics to multidimensional peptide and protein separations. For example, Rocklin and co-workers [40] have demonstrated 2-D separation of peptide mixtures in a microfluidic device using MEKC and CZE as the first and second dimensions, respectively. While MEKC and CZE are not perfectly orthogonal, they are perhaps the easiest to implement in a microchip with open channels and, when combined, still provide high peak capacity. In this example, a mixture of peptides is first separated by MEKC and then the effluent is sampled every 4 sec with a 0.3 sec injection to perform CZE in the second dimension, with total analysis times of less than 10 min for tryptic peptides. Approximately 10% of the sample injected into the first dimension is sampled by the second dimension and the peak capacity of the two-dimensional separations was estimated to be in the 500-1000 range. In a similar approach, Gottschlich et al. [41] reported on combining open-tubular reversed-phase chromatography with CZE for two-dimensional separation of peptides. A number of groups have also succeeded, at least partially, in adapting variants of 2D gel electrophoresis to microchips. True miniaturization of 2D gel electrophoresis has not been achieved yet because of the difficulties involved in the transfer of analytes from the first dimension to the second. This results mostly because of the incompatibility of the buffer systems used in the two separations - IEF needs highly-controlled mixture of ampholytes with low ionic conductivity while SDS-PAGE uses a highly conductive buffer. Leakage of SDS into IEF channel can completely ruin the IEF separation. Herr et al. [42] first demonstrated the integration between liquid-phase IEF and free solution CE by sequentially transferring samples


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from the IEF channel to an orthogonal CE channel. Once transferred to the CE channel, samples can be further separated based on their electrophoretic mobilities. Meanwhile, Wang et al. [43] implemented softlithography microfluidic valves to couple carrier ampholyte IEF with either CGE or CZE in a PDMS microchip. By active microvalve control, IEF focused samples can be isolated and selectively transferred to the second dimension separation channel. Further success in this area includes a plastic microfluidic network developed by Li et al. [44] and a multichannel differential gel electrophoresis (DIGE) platform presented by Emrich et al. [45]. Li et al. coupled nonnative IEF with SDS gel electrophoresis in a multi-channel platform. By controlling the channel resistance, sieving gel solution was first loaded into multiple separation channels. To increase the peak capacity (~1700), SDS solution was introduced between the first dimension IEF and second dimension gel electrophoresis to form SDS complexes [44]. Emrich et al. described a microfluidic platform integrating liquid-phase IEF and the second dimension CE with a fluid barrier created by passive valve structures (shallow etches). Cell lysate from 440,000 E. coli cells were successfully analyzed by the micro-DIGE analyzer as shown in Figure 5.5 [45].

Fig. 5.5 Schematic of the microchip to perform two-dimensional gel electrophoresis. The chip comprises an arced, 3.75 cm long horizontal channel for first dimension isoelectric focusing (IEF) that is punctuated with 20 6.8 cm long vertical channels through which focused proteins are separated in the second dimension by native gel electrophoresis. Reprinted with permission from [45]. Copyright (2007) American Chemical Society.

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5.3.3 Chips integrated with mass spectrometry Recent advances in speed, sensitivity and throughput of mass spectrometers has made them the detector of choice for many proteomic applications. The mass spectrometer as a detector offers many advantages including label-free detection and identification of a large number of peptides (and proteins) in a single run. Because of its high peak capacity, multidimensional liquid chromatography interfaced with mass spectromtery has become the technique of choice for genome-wide identification of proteins. Integration of microfluidic chip-based separations (e.g, chromatography) with mass spectrometry is attractive for a number of reasons including 1) ability to analyze smaller samples, 2) improved speed of analysis, and 3) facile integration of steps such as enrichment, separation and electrospray. The microchip interface to mass spectrometry has been achieved by a number of methods including creating a guided channel to the nanoelectrospray emitter using butted capillary [46], the double-etching procedure [47], polymer casting [48], and microdrilling [49-50]. One commercial product that allows integration of chip to a mass spectrometer is the Agilent's HPLC-Chip technology (www.agilent.com). The HPLC-Chip integrates sample preparation, separation, and electrospray tip on a single chip. The reusable chip is machined out of polyimide and contains a channel that serves as both separation column and electrospray nozzle. On-chip integration of sample, enrichment, and electrospray significantly reduces the number of fittings, connections, valves, and tubing required for conventional HPLC-MS and leads to higher-resolution separation and a more sensitive detection of peptides and other analytes.

5.4. Future Directions Microfluidic chips have made a significant impact in the field of protein analysis research and will continue to be used because of the numerous advantages they offer namely, faster, better, and cheaper analysis. Many conventional work-horse techniques such as zone electrophoresis, gel electrophoresis, isoelectric focusing, and limited forms of chromatography have been adapted to microchip format and the list will continue to grow. The biggest power of microfluidic chips, the ability to perform integrated and parallel operations, has not been realized yet. It is anticipated that in the future, progressively the focus will be on developing chips that integrate multiple functions and go towards realizing “sample-to-answer” de-


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vices. To date, focus has been on adapting single analysis techniques to a chip. In future, emphasis will shift to multi-dimensional analysis of proteins in a single chip and interfacing those separations with detectors such as mass spectrometers. While some successes have been achieved in integrating microfluidic chips to mass spectrometers, a lot of work needs to be done to achieve interfaces that are reproducible and easy to use. One of the problems has been that current mass spectrometers do not have the sensitivity to analyze the minute amounts that are typically analyzed in chips. With the development of more sensitive mass spectrometers and concomitant improvement in chip-based protein analysis and interfaces, it should be possible to have microfluidic devices that can be connected to any mass spectrometer.

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13. Yao S, Anex DS, Caldwell WB, Arnold DW, Smith KB, Schultz PG (1999) SDS capillary gel electrophoresis of proteins in microfabricated channels. Proc Natl Acad Sci USA 96:5372-5377. 14. Bousse L, Mouradian S, Minalla A, Yee H, Williams K, Dubrow R (2001) Protein Sizing on a Microchip. Anal Chem 73:1207-1212. 15. Han J, Singh AK (2004) Rapid protein separations in ultra-short microchannels: microchip sodium dodecyl sulfate-polyacrylamide gel electrophoresis and isoelectric focusing. J Chromatogr A 1049:205-9. 16. Wu J, Pawliszyn J (1992) Application of capillary isoelectric focusing with universal concentration gradient detector to the analysis of protein samples. J Chromatogr 608:121-30. 17. Herr AE, Molho JI, Drouvalakis KA, Mikkelsen JC, Utz PJ, Santiago JG, Kenny TW (2003) On-chip coupling of isoelectric focusing and free solution electrophoresis for multidimensional separations. Anal Chem 75:1180-7. 18. Tan W, Fan ZH, Qiu CX, Ricco AJ, Gibbons I (2003) Miniaturized capillary isoelectric focusing in plastic microfluidic devices. Electrophoresis 23:363845. 19. Li Y, Buch JS, Rosenberger F, DeVoe DL, Lee CS (2004) Integration of isoelectric focusing with parallel sodium dodecyl sulfate gel electrophoresis for multidimensional protein separations in a plastic microfluidic network. Anal Chem 76:742-8. 20. Xu Y, Zhang CX, Janasek D, Manz A (2003) Sub-second isoelectric focusing in free flow using a microfluidic device. Lab Chip 3:224-7. 21. Sommer GJ, Singh AK, Hatch AV (2008) On-Chip Isoelectric Focusing Using Photopolymerized Immobilized pH Gradients. Anal Chem 80:3327-33. 22. Reichmuth DS, Shepodd TJ, Kirby BJ (2005) Microchip HPLC of Peptides and Proteins Anal Chem 77:2997–3000. 23. Terabe S, Otsuka K, Ichikawa K, Tsuchiya A, Ando T (1984) Electrokinetic separations with micellar solutions and open-tubular capillaries. Trends Anal Chem 56:111-113. 24. Nota G, Marino G, Buonocore V, Ballio A (1970) Liquid-solid Chromatography with Open Glass Capillary columns. J Chromatogr 46:103. 25. Knox JH, Gilbert MT (1979) Kinetic Optimisation of Straight Open-tubular Liquid Chromatography. J Chromatogr 186:405-418. 26. Rohr T, Yu C, Davey MH, Svec F, Fréchet JM (2001) Porous polymer monoliths: simple and efficient mixers prepared by direct polymerization in the channels of microfluidic chips. Electrophoresis 22:3959-67. 27. Throckmorton DJ, Shepodd TJ, Singh AK (2002) Electrochromatography in microchips: reversed-phase separation of peptides and amino acids using photopatterned rigid polymer monoliths. Anal Chem 74:784-9. 28. Bedair M, Oleschuk RD (2006) Lectin affinity chromatography using porous polymer monolith assisted nanoelectrospray MS/MS. Analyst 131:1316-21. 29. Hatch AV, Herr AE, Throckmorton DJ, Brennan JS, Singh AK (2006) Integrated preconcentration SDS-PAGE of proteins in microchips using photopatterned cross-linked polyacrylamide gels. Anal Chem 78:4976-84.


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Chapter 6 Microfluidic Systems for Cellular Applications

H. Tavana, Y.-K. Chung, C.-H. Kuo and S. Takayama Department of Biomedical Engineering, University of Michigan, Ann Arbor, MI 48109 Correspondence should be addressed to: Prof. S. Takayama ([email protected])

Keywords: Microfluidics, Cell Isolation, Cell Culture, Cell Analysis, Cellular Microenvironment, Cell Signaling

Abstract Cells are the basic building units of the vast diversity of living organisms. Study of cells and their functions is of great importance in pure and applied sciences and spans a wide range of areas including cell biology, human physiology, and tissue engineering. Much of our knowledge of intricate cellular behavior and functions is due to various techniques that allow in vitro manipulation of cells. Handling of cells outside the body often involves the three main steps of isolation, culture, and analysis. Existing traditional methods have facilitated these processes and revealed a wealth of information. Nevertheless, recent advances in our understanding of various aspects of cellular systems point to the inadequacy of conventional techniques. For example, conventional cell culture systems do not mimic intricate in vivo cellular microenvironments and as a result, many cellular phenotypes are lost. The emergence of microfluidic technology in the past

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decade and its compatibility with all three cell handling processes has given a new spin to study of cells in various contexts. Microfluidic settings for the isolation, culture, and analysis of cells offer advantages over their conventional counterparts by providing a more physiologic microenvironment for cells, enabling various biologically important culture conditions (e.g. static and dynamic), and being economically sounder. The commercialization of microfluidic cell-based platforms also shows a promising trend and new products steadily find their way into the market.

6.1 Introduction Cells are the simplest structural units into which a complex multicellular organism can be divided and still retain the functions characteristic of life. Cells of the body arrange themselves in various combinations to form a hierarchy of organized structures ranging from tissues, to organs, to organ systems, and finally to a total organism. The response of an organism to disease, injury, or therapy is therefore a collective response of its cells. For decades, in vitro cell culture has served as a gateway to explore various aspects of cellular biology. The ease of introducing systemic alterations to the environment of cells and the possibility of modulating the phenotype of cells has prompted an unprecedented expansion of in vitro systems with the aim of designing cell-based assays, sensors, and substituents of native tissues. Engineered in vitro constructs have also served as models for quantitative studies of cell and tissue response to genetic alterations, drugs, hypoxia, and mechanical stimuli. Studies of cells often involve three key steps: separation of a cell type of interest from a heterogeneous population, culture of the isolated cells, and performing biochemical analyses on them. Standard procedures exist for each of these steps; however, they entail a number of limitations that are briefly discussed below. Conventional methods for separation of a cell type of interest from a heterogeneous population are based on differences in physicochemical properties of different cell types including size, density, and electric charge [14]. However, these properties vary only slightly from one cell type to another, making cell separation processes inefficient. Immunologic techniques, such as magnetic cell sorting and fluorescence-based flow cytometry, involve selectively attaching magnetic or fluorescent particles to cells

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and yield higher efficiencies [1,5]. Nevertheless, these separation methods require preparatory and incubation time periods, actuation and detection systems, and post-processing of cells such as the removal of labeling. Overall, traditional approaches for cell separation are costly, timeconsuming, mechanically complex, and require trained personnel for operation and maintenance. They can also be lethally or sub-lethally damaging to cells. The history of cell culture dates back to early twentieth century. The original drive for the development of cell cultures was to investigate various physiological events such as nerve development [6]. It was in the 1940s and 1950s that animal cell culture became a routine laboratory technique. Historically, in vitro culture of animal cells has been conducted on simple, homogeneous surfaces such as those of Petri dishes. It is well known that the cells in these in vitro environments often do not express the properties characteristic of the same cell type in vivo. The problem stems from the lack of physiologic cellular microenvironment in these in vitro culture systems. Inside the body, cells reside in a tissue specific microenvironment and are subject to multiple cues that vary in time and space, including gradients of various soluble factors (hormones, growth factors, nutrients, and inorganic ions), biochemical and mechanical interactions with the extracellular matrix (ECM), and direct cell-cell contacts. Conventional culture systems do not provide such a physiologic environment for cells and as a result, only limited information can be gained about the phenotype of cells grown in these systems. The following example elucidates this point. In a cell culture dish, the volume of culture media above cells is significantly greater than the volume taken up by the monolayer of cultured cells. Continuous secretion of signal molecules by cells, which regulates their own functions as well as functions of their neighboring cells, causes spatial variability in the concentration of solutes in the culture media. This concentration gradient initiates convective mass transport and causes rapid distribution of the molecules over the entire volume of the media. As a result, autocrine and paracrine signaling is, at least temporarily, impaired [7] (Note: autocrine and paracrine signalings refer to the binding of a signal molecule to receptors on the surface of the same cell that secrets it and the neighboring cells, respectively, to initiate a cascade of intracellular and intercellular events). It is not difficult to think of other examples that highlight differences between the environmental conditions of a cell in vivo and in vitro. From the viewpoint of the pharmaceutical industry, the microenvironment disparity has significantly hindered the development of new drugs based on tests conducted with conventional cell-based assays to screen drug safety and efficacy.

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Macroscale analytical techniques for biochemical analyses of cells also have several major drawbacks: These techniques mainly rely on large and expensive bench-top equipment, which makes them incompatible with point-of-need applications. They often require a large number of cells, which is not always available (such as in the study of precious cells and cell samples from human or animal models). In the case of single cell analysis, existing macroscale analytical techniques are slow and only a small number of cells can be analyzed per day [8]. The manually intensive nature of these methods is a major reason for their low throughput. Enhanced fabrication capabilities of the microelectronics industry in the early 1980s led to the advent of microelectromechanical systems (MEMS) that integrate mechanical components with electronics to create functional devices [9]. The technology was extended later to the manipulation of fluids inside micrometer scale channels and wells [10,11] and thus, the field of microfluidics emerged. A major source of motivation for microfluidics came from developments in different areas of biology such as molecular biology that required analytical methods with high throughputs and resolution not achievable previously with conventional methods. The commensurability of the linear dimensions of eukaryotic cells (10-100 µm) with microfluidic settings opened up the opportunity to use this technology for cellular applications. Microfluidics not only enables carrying out the three main processes of separation, culture, and analysis of cells, it also enjoys many inherent advantages over the macroscale cell handling counterparts. The benefits of microfluidic technology for cellular applications are discussed below from physiological, biological, and economical perspectives.

6.1.1 Physiological advantages A glance at the human anatomy reveals a large number of networks that are microfluidic in nature. These networks exist at different hierarchical levels, from system level such as the nervous, the lymphatic, and the cardiovascular systems to the functional units of various organs such as nephrons in the kidney, the network of alveolar ducts and sacs and the corresponding blood capillaries in the lung, etc. (Fig. 6.1). It therefore makes intuitive sense to mimic such in vivo structures with appropriate microfluidic devices for in vitro cell-based studies. Microfluidic technology also makes it possible to recreate tissues of different organs on a single chip by culturing various types of cells at different sections of an interconnected network of channels and chambers [12]. Such “body-on-a-chip” (BOC)


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platforms would allow communication between distant cells through endocrine signaling, revealing information that could not be learned from conventional cultures. Endocrine signaling plays an important role in regulating the response of the human body to various pharmaceuticals and chemicals. Therefore, functional BOCs can provide realistic and physiologic platforms for drug testing, reduce the need for animal-based tests and lower the cost of developing new pharmaceuticals. A more distant prospect of BOCs is customizing the treatment of patients with severe disorders. Cell samples from individual patients can be cultured in a chip and responses to particular drugs can be assessed to determine the most effective treatment. (a)

(b)

Terminal Bronchiole Branch of Pulmonary Vein Branch of Pulmonary Artery

Smooth Muscle

Respiratory Bronchiole

Alveoli Capillary

300 µm

Fig. 6.1 Many systems and functional units of the body are microfluidic in nature such as (a) the vessels of the lymphatic and cardiovascular systems in neck and head (Image reproduced, with permission, from 3D4Medical.com/Getty Images [13]) and (b) the respiratory bronchioles with clusters of alveoli and blood vessels (Image reproduced, from ref. [14] with permission from The McGraw-Hill Companies (©2006)

6.1.2 Biological advantages Microfluidic cellular systems are valuable tools from a biological standpoint too. First, both static and dynamic culture conditions can be realized in microfluidic devices. Given that convection is negligible in microchannels that are not perfused and diffusion is the operative mode of molecular transport, static cell cultures preserve autocrine and paracrine factors in the

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proximity of cells. The formation of gradients of soluble factors in the channels is physiologically significant as it is a key regulator of the fate and the phenotype of various cells. Dynamic cell culture is an even more appealing feature of microfluidic systems since it enables the recreation of various physiologically-relevant processes. For example by controlling the flow rate in microchannels, cell-cell interactions through autocrine, paracrine signaling, and even endocrine effects can be systematically manipulated. With dynamic cultures, it is also possible to mimic a range of in vivo processes directed by mechanical stimulation of cells, such as the modification of morphology of endothelial cells under shear stress [15], rolling of leukocytes along the luminal surface of blood vessels under blood flow [16], and injury of small airway cells due to fluid mechanical stresses generated by propagation of liquid plugs during reopening of closed pulmonary airways [17]. Second, materials with tunable properties might be used for the fabrication of microfluidic devices and different geometrical features can be incorporated inside microchannels to investigate various aspects of cell-ECM interactions. For example, the effect of ECM elasticity on the fate of embryonic stem cells was studied by tailoring surface properties of ECM [18]. Some aspects of cell-ECM adhesion characteristics were revealed through measurements of forces exerted by cells on an array of microposts [19], and 3D scaffolds were incorporated in microchannels to provide cells with a more in vivo-like ECM environment [20,21]. Third, fluid flow in microchannels is almost always laminar with typical Reynolds numbers of less than one. There is no appreciable mixing between streams of fluids other than by diffusion. Laminar flow is one of the key features of microfluidic systems and is exploited for a wide range of on-chip applications including subcellular positioning of small molecules [22], protein fractionation [23], and rapid immunoassays [24]. Overall, laminar flow provides a gentle environment for separation, culture, and analysis of cells. Fourth, there is a growing emphasis in molecular biology on single cell analysis to understand phenotypic heterogeneity among cells in a population. Microfluidic systems are advantageous since they can enhance the throughput of single cell analysis and integrate various cell handling and processing steps [25].


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6.1.3 Economical advantages The application of microtechnology to cell biology can realize significant economical advantages over conventional cell handling processes. Microfluidic devices have small volumes and large surface area-to-volume ratios. As a result, only small volumes of reagents (several nanoliters) are consumed and fluidic reaction times are significantly shortened. Minute reagent volumes also reduce the amount of chemical waste produced. The process of device fabrication can be straightforward and commonly-used polymer materials for fabrication, e.g. poly(dimethylsiloxane) (PDMS), are relatively inexpensive (e.g. compared to silicon microdevices). Furthermore, high throughput microfluidic platforms can be realized by including many copies of a design on a single chip to simultaneously test the effect of different samples, say of a drug, on cells. It is also possible to integrate all three steps of cell separation, culture, and analysis into a single chip. Integrated devices can be complex and challenging to operate but when successful, reduce processing cost and time. The small footprints and low power requirements also make them suitable for portable, point-of-need applications. The discussion above outlined the shortcomings of conventional techniques for cell studies and highlighted microfluidic technology as a useful alternative. In the next section, although far from exhaustive, a selection of novel microfluidic platforms developed for separation, culture, and analysis of cells is presented. Then, commercialized microfluidic devices for cellular applications will be briefly reviewed. Finally, the chapter will conclude with future prospects of microfluidic cell systems.

6.2 Microfluidic Technology for Cellular Applications 6.2.1 Microfluidic cell isolation/separation Separation and isolation of cells is a key process for a variety of applications including: Fractionating whole blood samples to isolate white and red blood cells, deriving helper T lymphocytes to monitor HIV treatments, capture of rare circulating tumor cells to identify key biological determinants of blood-borne metastases [26], and deriving a stem cell line from mature tissues or liposuction aspirates. Commercially available cell separators often have several limitations such as large foot-print, the need for expensive lasers and detectors as well as trained operators.

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Microfluidic systems are low-cost and easy-to-use alternatives. Downscaling cell separation systems to dimensions comparable to that of cells and harnessing unique fluidic properties at the micrometer scale results in increased speed, reduced sample consumption, smaller foot-print, and higher throughput. Separation mechanisms based on dielectrophoresis, capillary electrophoresis, acoustic forces, and optical traps can all be incorporated into microfluidic devices [27-34] and optical components can be integrated into these platforms. Below, a number of microfluidic cell separation methods are described. The selection highlights the flexibility of microfluidic systems to employ physical and biological principles for separation/isolation of different cell types including precious cells. Simplicity of fabrication, small reagent volumes, gentle separation, fast separation time, and user-friendliness are common features of these settings. 6.2.1.1 Microfabricated fluorescence-activated cell sorter (µFACS)

Conventional fluorescence-activated cell sorters (FACSs) are widely used in clinical medicine and biological sciences. FACSs are remarkably efficient, however they are costly, mechanically complex, difficult to sterilize, and require large sample volumes as well as trained personnel for operation/maintenance. Inexpensive devices that rapidly sort cells and particles would facilitate screening of cell populations and combinational chemistry libraries. Fu et al. developed an elastomeric microfabricated FACS (µFACS) based on electroosmotic flow [35]. The device resembled a T-junction with inlet, collection, and waste chambers. Two sorting algorithms were used: “forward” and “reverse”. In the “forward” method, cells flowed in a buffer solution from the input channel to the waste channel. If the fluorescent intensity of a cell was above a preset threshold, the voltages were temporarily changed to divert the cell to the collection channel. The “reverse” method was used to identify rare cells. Cells were moved at a high speed from the input to the waste. The flow was stopped when a cell of interest was detected. By the stoppage time of the flow, the cell was past the junction toward the waste channel. The system was then run backward at a slow speed from the waste to the input, and the cell was switched to the collection channel when it passed through the detection region. The device was used to sort wild E. coli cells from GFP-expressing E. coli cells. An enrichment of 80- to 96-fold and a sample throughput of 10-20 cells/sec was


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achieved. Later, an optimized device from this group gave a two fold increase in the throughput [36]. However, this was still several orders of magnitude lower than that of conventional FACS, i.e. 10000-20000 cells/sec. To reduce the efficiency gap between conventional and microfabricated FACSs, Wolff et al. designed a pressure-driven µFACS silicon chip with three inlets and two outlets [37]. A sample of chicken red blood cells (CRBCs) and fluorescent latex beads was introduced from a middle inlet and sheathed by buffer streams from two sides. The fluorescent beads were excited near the junction of collecting and waste channels. The fluorescence signal was detected by a photomultiplier tube and the system activated a valve to force cells of interest to the collecting channel. A 100-fold enrichment of fluorescent beads was achieved at a throughput of 12000 cells/sec, i.e. 1000 times higher than that of previous µFACS. In a separate design, a different configuration was used to generate hydrodynamic focused sample flow. The sample was introduced into the buffer stream through a “chimney” like structure, being carried downstream the channel like a thin wisp of smoke from a chimney. This ensured lower variations in the fluorescent signal, which is a common problem in devices with hydrodynamic focusing of sample using side sheath flows. Other interesting features of this chip were the integration of a chamber for holding and culturing the sorted cells and optics for the detection of cells on chip. Such devices are promising signs for developing fully integrated functional microfluidic platforms for cell sorting, culturing, and analysis. Wang et al. described a µFACS that utilizes optical forces to sort mammalian cells [38]. A sample of Hela cells stably transfected with histone-GFP mixed with non-expressing parental Hela cells flowed from inlet and were sheathed to a narrow stream by hydrodynamic focusing using side flows. Cells in the focused flow first passed through an analysis section and then through an optical switching region of the device. Flow to the two output channels was asymmetrically biased and with no optical beam present, all cells moved to the waste channel. When a target cell was detected, the optical switch was activated and a focused laser spot deflected the cell to the target output channel. Because this switch operates by displacing the cell within the laminar flow rather than by making a transient change to the fluid flow, switch rates can be high. Throughputs of up to 106 cells/sec, recovery rates of >85%, and collection well purities of up to ~83-99% were obtained. Cell viability was evaluated by examining the expression levels of two cellular stress indicator genes HSPA6 and FOS. The results

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showed no increase in the expression levels of these genes, an indication of viable cell populations. µFACSs have been optimized to yield reasonably high enrichments and throughputs. These settings are particularly useful for sorting of precious cells and applications where only a small number of cells is available, e.g. 100-100,000 cells, which cannot be handled with conventional flow cytometers. In addition, µFACSs offer many other advantages including consumption of small volumes of reagents, low cost, easy device sterilization, minimum sample carryover, and potential to integrate various other functional units to realize a fully self-contained lab-on-a-chip platform. 6.2.1.2 Microfluidic magnetic cell separation

Cells can be isolated under the influence of a magnetic field. Magnetic cell separation (MCS) is divided into two classes [39]: (i) separation using native magnetic susceptibility of cells and (ii) separation using antibodycoated magnetic beads. The former technique relies on the property of certain cells such as deoxygenated red blood cells to contain paramagnetic material [40-42]. Paramagnetic cells can be separated from the cell suspension under a magnetic field. Microfluidic devices that use this property to separate a cell type of interest contain a small ferromagnetic wire loosely packed into a region near the poles of a large magnet. The small wires generate high magnetic field gradients when magnetized and trap the paramagnetic cells. Han et al. demonstrated continuous separation of red blood cells in a microfluidic device by passing whole blood around a 50 µm magnetized nickel stripe [43]. Cell separation can also be realized through specific binding of antibody-coated magnetic beads with antigens expressed on a cell surface [44,45]. A number of microfluidic platforms have been described for the separation of magnetic beads [46-48] and cells [49-51] using this technique . For example, Inglis et al. incorporated magnetic strips into a silicon substrate and magnetized them by applying an external field [49]. The strips were placed at an angle to the direction of fluid flow carrying narrow streams of cells over the stripes. Magnetically labeled cells were attracted to the strips, whereas unlabeled cells did not interact with the strips and followed the direction of fluid flow. Separation of leukocytes from whole human blood was demonstrated. MCS is an inexpensive method for the separation and enrichment of different cell types including tumor cells. However, preventing permanent adhesion of the magnetically labeled cells to the strips is a great challenge.


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Normally, a significant number of magnetically labeled cells (~50%) either permanently stick to the nickel strips or are not sufficiently attracted to the strips to be separated [39]. 6.2.1.3 Transient cell-ligand adhesion

Cells may be isolated by recruiting natural, physiological process of leukocyte adhesion to blood vessel walls at the sites of injury or inflammation. For example during an inflammatory response, glycoproteins of white blood cells (WBCs) are recognized by cell adhesion molecules, including the selectin family, on the surface of endothelial cells (ECs) lining blood vessels at the inflammation site. This recognition causes WBCs to adhere to the blood vessels. This transient adhesion between WBCs and surface ligands retards movement of the cells. Due to a continuous dissociation/formation of bonds between proteins on the surface of WBCs and ECs under shear from the blood flow, WBCs roll slowly along the blood vessels luminal surface and eventually, migrate to the inflamed tissue. Chang et al. used this principle to capture and concentrate cells in microfluidic devices [16]. Channels contained arrays of either square or offset micropillars with spacing larger than the size of the cells to allow for free motion of the cells. Channels were treated with human E-selectin IgG as the adhesion protein. A suspension of HL-60 cells bearing ligands for Eselectin was flowed at 1 µl/min. In the first minute of the experiment, channels with offset pillars captured about 95% of the cells over a distance of 700 µm whereas the capture efficiency of the square array design was about 72%. With increasing the flow time, the efficiency of the square geometry improved to 85-94%. Both designs were capable of enriching cells several hundred folds from the original concentration. This technique is potentially applicable to chromatographic fractionation of adherent cells because different cell types have different affinities for a given adhesion molecule. Separation of HL-60 and U-937 myeloid cells was demonstrated, albeit with low resolution. A similar strategy was applied by Nagrath et al. to separate circulating tumor cells (CTCs) from peripheral whole blood samples in a single step without a need for pre-dilution, pre-labeling or other sample processing steps [26]. CTCs are rare and comprise as few as one cell per 109 haematologic cells in the blood of patients with metastatic cancer. The separation is mediated by the interaction of target CTCs with anti-epithelial cell adhesion molecule (EpCAM)-coated array of microposts incorporated in the channels. Overexpression of EpCAM in the carcinomas of lung, breast,

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and prostate provides the specificity for CTCs capture from the nonfractionated blood. CTCs were identified in 99% of the blood samples donated by cancer patients. The purity of CTCs capture varied between ~50% to ~70% for cancers of different origin and a capture specificity of 100% was achieved. 6.2.1.4 Laminar flow

Approximately 10% of couples have infertility problems [52]. The most advanced treatment for male-related infertility is an in vitro fertilization (IVF) technique known as intracytoplasmic sperm injection (ICSI) [53], where an oocyte is fertilized by the direct injection of a sperm. To maximize the rate of successful pregnancy and birth, it is important to select the most viable sperm. Although ICSI has significantly reduced the number of viable sperm required for fertilization, it bypasses all sperm selection processes. Due to the limitations of current techniques such as centrifugation and swim-up processes [54] for the isolation and selection of viable sperms, doctors frequently resort to hand sorting through dead sperm and debris, a procedure that can take hours in some cases. Developing novel processes to isolate the most viable sperm would greatly benefit IVF processes in clinical settings. Cho et al. described a PDMS microscale integrated sperm sorter (MISS) that separates motile sperm from non-motile sperm and cellular debris [55] (see Fig. 6.2). Horizontally oriented reservoirs with different heights at inlets and outlets provided a passively driven gravity pumping system and facilitated liquid movement in the channels. Separation in this microfluidic system relied on the existence of multiple parallel laminar streams with no mixing at the interface between them. Non-motile human sperm, ~60 µm in length, and particles diffused slowly and remained within their initial streamlines. In contrast, motile sperm were very mobile and swam at ~20 µm/sec and distributed randomly within the 500 µm-width channel. A bifurcation at the end of the separation channel was used to collect motile sperm that deviated from its inlet streamlines (Fig. 6.2). The purity of motile sperm after sorting was nearly 100%, regardless of motile sperm purity prior to the sorting. The yield of the MISS, i.e. the ratio of the number of collected motile sperm to the total number of motile sperm at the inlet reservoir was ~40%, which is comparable to the recovery rates of conventional sperm sorting methods [54,56]. In principle, this separation mechanism is similar to that of “H-filter” where rapidly diffusing small molecules exit through a different outlet from large molecules and particles that diffuse slowly [57,58]. The main difference is that the MISS takes


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advantage of active movement of cells whereas the H-filter relies on the passive diffusion of molecules.

Fig. 6.2 Three-dimensional illustration of the microfluidic channel design with horizontally oriented fluid reservoirs and separation of motile sperm from nonmotile sperm (Reprinted with permission from [55] Copyright (2003) American Chemical Society.).

The MISS has several advantages over existing techniques: (i) it is a selfcontained functional microdevice that manipulates cells without a need for electronics and external power sources, (ii) it offers an efficient way for the isolation of sperm from samples with few motile sperm that are difficult or impossible to process using conventional methods, (iii) it facilitates a mild biomimetic process and avoids centrifugation and sample compaction that are known to cause sublethal damage to sperm [54,59], (iv) it is small, portable, easy to use, disposable, and inexpensive. The MISS is promising for use in large scales in clinical settings and opens up the possibility of making self-contained bioassays for convenient at-home screening of male infertility.

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6.2.1.5 Deterministic hydrodynamic

A microfluidic device was described by Davis et al. that separates WBCs and red blood cells (RBCs) from whole blood and fractionates different WBCs (lymphocyte, monocyte, granulocyte, T cell, B cell) according to their size [60]. WBCs are more or less spherical and range from 5 to 20 µm in diameter, whereas RBCs are disk-shaped with 8 µm in diameter and have a 2 µm thickness. The principle of operation is as follows: A cell suspension flows through an array of microposts incorporated in the channel. Each row of posts is slightly offset laterally with respect to the row immediately above it. Particles smaller than a critical hydrodynamic diameter (Dc) follow streamlines cyclically through the gaps, moving in an average downward flow direction. Particles larger than Dc do not fit into the first streamline and bump into a neighboring streamline at each post. Thus they move at an angle with respect to the direction of fluid flow, determined by the ratio of post offset to the row-to-row spacing. Fig. 6.3 illustrates the design. Whole blood and a buffer solution entered into different channels at the top of the device and maintained a vertical laminar flow throughout the device. The active region consists of 13 consecutive regions with various Dc of 3 to 9 µm. Cells smaller than 3 µm moved straight down, those larger than 9 µm bumped at all regions of the device, and cells of intermediate size started by bumping in the upper regions of the device and eventually moved straight after entering a region with a Dc larger than the cell size. The bumped cells were diluted into the buffer solution. The device had three separate output channels to collect cells of slightly differing hydrodynamic radii. Cells behaving as objects larger than the maximum Dc for all 13 regions (>9 µm diameter) were thus displaced from the left edge to the right edge, exiting from channel 3, whereas those with a diameter of less than 3 µm passed straight down the device and exited from channel 1. The analysis of collected cells showed that more than 99% of all WBCs were displaced into channels 2 and 3 and the distribution was dependent on the cell size. For example, the ratio of lymphocytes in channel 2 to channel 3 was ~100 to 1. The total number of RBCs at the three outlets was also determined as 3×106 with 99% of the RBCs exiting from channel 1.


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Fig. 6.3 Microfluidic device to separate WBCs from RBCs and platelets. Arrows indicate the vertical and bumping directions (Image reproduced, with permission, from ref. [60] Copyright (2006) National Academy of Sciences, U.S.A.).

6.2.1.6 Free flow acoustophoresis

Acoustic forces are widely used to separate suspended particles from their medium or from other particles [61,62] as well as to trap particles [63,64]. Acoustic separation and trapping systems are gentle to biological systems and also easy to operate. Petersson et al. described a microfluidic platform that utilizes the combination of laminar flow and acoustic forces to separate particles and cells based on their size and density in an ultrasonic wave field [32]. The technique is called free flow acoustophoresis (FFA). Suspended particles/cells entered a 350 µm-wide channel through two side inlets. The medium without suspended particles flowed through the center inlet into the main channel. An acoustic force field perpendicular to the flow direction was generated between the side walls of the channel using a piezo ceramic actuator. As the particles/cells moved along the flow path, the acoustic force displaced them toward the center of the channel. By balancing the flow rate, acoustic force, and particle mixture, a particle gradient developed across the channel. Due to the laminar flow in the channel, this gradient was further fractionated down the channel into several outlets. Fractionation of suspension of particles/cells with fairly similar densities was more challenging and required the addition of small amounts of cesium chloride to the medium to manipulate its density. The strategy was proved useful to fractionate a suspension of RBCs and platelets containing 1% by volume from each cell type in a device with two active outlets. 92% of the RBCs and 99% of the platelets were directed to the first and the sec-

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ond outlets, respectively. A more complex cell suspension, i.e. Buffy coat, was also fractionated by FFA. Blood component therapy for transfusion, i.e. the use of pure RBCs, platelets, and leukocytes, as well as in modern transfusion therapy based on leukocyte-depleted platelets may benefit from such microdevices.

6.2.2 Microfluidic cell culture In vitro culture of animal cells has a wide range of applications. For example: to study the normal physiology or biochemistry of cells such as cell metabolism, to investigate the sequential or parallel combination of different cell types to generate artificial tissues, to evaluate the safety and efficacy of new pharmaceuticals, to synthesize valuable biologicals such as therapeutic proteins from large scale cell cultures, etc. Standard culture systems that are often performed on 2D treated plastic surfaces do not mimic the in vivo microenvironment of cells: oxygen tension is too high, the concentration of soluble factors is abnormally high, cell-cell interactions are unorganized, and 3D cues specific of the native tissue are absent [25]. Microfluidic technology provides the opportunity to create a physiologic cellular microenvironment by regulating the transport of nutrients and other biochemicals in microchannels in a spatio-temporal manner and engineering the ECM using different materials and geometries to mimic complex ECM in vivo and to provide cells with appropriate extracellular cues. The application of microfluidics to the culture of mammalian cells is discussed below in the context of a few examples. These studies demonstrate the feasibility of creating physiologic tissue models through controlled manipulation of the microenvironment of cells, an achievement that could not be realized previously with conventional dishbased cultures. 6.2.2.1 Recreation of pulmonary airways on a chip

The branching network of pulmonary airways is lined with a viscous liquid film secreted by airway epithelial cells. The primary function of airways is directing airflow to the sites of gas exchange with the blood. Many respiratory diseases such as chronic bronchitis, emphysema, and asthma are accompanied by overproduction of mucus in airways and often, by dysfunction of pulmonary surfactant. As a result, the free surface of the thick mucus-containing liquid film grows over time and eventually forms a liquid plug across the airway lumen. The liquid plug obstructs airflow, result-


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ing in poor ventilation of the lung and insufficient blood oxygenation. Upon inhalation, the liquid plug propagates along the airway and eventually ruptures, reopening the occluded airways (Fig. 6.4a). This process is known as “airway closure and reopening” [65,66]. Movement of liquid plugs and reopening of occluded airways is known to generate large fluid mechanical stresses [67], which are suspected to damage small airway epithelial cells (SAECs). Nevertheless, the complexity of pulmonary fluid flows and the small size of airways (several hundred micronmeters) have hindered experimental study of the cellular level injury in vitro. Huh et al. developed a three-dimensional microfluidic device that closely mimics the microenvironment of airway epithelial cells in vivo [17]. The microfluidic airway system consists of two PDMS compartments separated by a polyester porous membrane (Fig. 6.4b). The upper and lower compartments represent the apical and basal chambers of airway epithelium, respectively. Primary human SAECs were seeded into the upper chamber and cultured on the membrane, which mimics the basement membrane. To promote the growth of the cells, both chambers were perfused with culture media for about one week. Once a confluent monolayer of cells was formed, the apical surface of the cells was exposed to an air-liquid interface to induce cellular differentiation. Maintaining the microfluidic culture system under this condition for about two weeks induced cellular differentiation, causing SAECs to express morphological and secretory phenotypes found in native airway tissues. Differentiation of SAECs was confirmed by immunoassay of Clara cell secretory protein as well as by immunohistochemical detection of tight junctions between epithelial cells. After three weeks of microfluidic culture, over 85% of cells were still alive. The microfluidic airway system was then integrated with a computerized liquid plug generator to study cellular damage during airway reopening. Preliminary results showed that plug propagation and rupture is deleterious to SAECs and cellular viability decreases significantly with increasing number of propagation and rupture events. The injurious effect of airway opening on lung epithelial cells was also reported by Bilek et al. using the propagation of a semi-infinite bubble in a narrow fluid-occluded channel lined with cells [68]. Such studies can enhance current understanding of cellular responses to complex mechanical forces of the pulmonary system and contribute to the design of strategies for preventing/treating lung injuries.

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(a)

(b)

Air flow

Rupture

Fig. 6.4 Schematic of (a) occluded pulmonary airways and the reopening process, and (b) micro-engineered compartmentalized biomimetic small airway system (Image reproduced, with permission, from ref. [17] Copyright (2007) National Academy of Sciences, U.S.A.)

6.2.2.2 Liver on a chip

Many drug candidates fail clinical trials either because they damage the liver directly or because liver metabolites are toxic [69]. The problem stems, at least in part, from the inadequacy of standard cell-based assays for screening drug safety and efficacy. Development of new drugs and the process of drug discovery would greatly benefit from in vitro cell culture systems that can maintain the physiological liver functions of hepatocytes. However, the development of reliable liver cell cultures as “biosensors” for drug toxicity is quite challenging mainly due to the difficulty in maintaining differentiated phenotypes over prolonged periods. Dordick and co-workers developed a 3D cell culture array for highthroughput toxicity screening of drug candidates and their metabolites against different cell types [70,71]. A microarray of MCF7 breast cancer cells encapsulated in a hydrogel matrix (collagen or alginate) was spotted onto a functionalized microscope glass slide and then was incubated in culture medium for six days. To evaluate the response of cells to different cytotoxic compounds, this slide was stamped against a microarray of various toxins immobilized on a second glass slide. Cells were grown for three days prior to performing the viability assay. Griffith and co-workers developed a microbioreactor for the in vitro culture of liver tissue under continuous perfusion [72]. The bioreactor is integrated with a 3D scaffold that plays the ECM role. The scaffold consists of


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a thin silicon sheet permeated with a regular array of microchannels from top to bottom. This scaffold sits on top of a microporous filter, which in turn is mechanically supported by a second scaffold. The reactor dimensions combined with the perfusate flow rates meet estimated values of cellular oxygen demands and provide shear stresses below physiological range. The channel walls were covered with a fluoropolymer during deep reactive ion etching (DRIE) of the silicon wafers and were subsequently coated with collagen I to enhance the adhesion and guide the morphogenesis of hepatocyte cells. Hepatocytes were seeded both in form of single cell suspension and pre-aggregated multicellular spheroids. In both cases, tissue-like structures formed inside the reactors. However, the microtissues formed from single cell suspension remained stable for four days only and then a progressive loss of structure was observed. On the other hand with hepatocyte spheroid seeding, the architecture and viability of tissues were preserved for up to two weeks and cultures exhibited cell-cell contact such as tight junctions and desmosomes, found in tissues in vivo [69,72]. Hepatotoxicity studies will benefit from the availability of such microengineered systems to investigate in vivo physiology and pathology in an in vitro environment. It has been observed that co-culture of hepatocytes with other cell types, including liver epithelial cells and Kupffer cells, prolongs the survival of cultured hepatocytes for up to two months and helps maintain liverspecific properties such as albumin secretion [73]. Furthermore, it is known that nonparenchymal cells participate in pathophysiological responses including drug toxicity and infection in conjunction with hepatocytes [72]. These findings emphasize that more physiological liver cell cultures representing a wide range of liver functions in vitro might only be realized by including appropriate homo- and heterotypic cell-cell interactions and possibly by considering various aspects of the liver microarchitecture in vivo. For example, Bhatia and co-workers described a 24-well culture system with microscale architecture fabricated using soft lithography for human liver cells [74]. Each well contained 37 colonies of hepatocytes on 500 µm diameter collagenous patterns that were surrounded by 3T3 fibroblasts. Unlike pure cultures that rapidly lost their morphological features and liver specific functions, the micropatterened co-cultures were stable and maintained liver specific functions for several weeks, as confirmed by different tests including albumin secretion, urea synthesis, and gene expression levels of liver-specific genes.

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6.2.2.3 Microfuidics for stem cell research

Stem cells (SCs) are capable of dividing extensively and developing into many different types of cells in the human body. SCs hold great promise for cell-based therapy of many diseases including heart, lung, and blood diseases, diabetes, Parkinson’s and Alzheimer’s diseases as well as the repair of injured tissues and organs [75]. The main challenge is to understand differentiation pathways of SCs into various cell types and control the behavior of SCs in culture. Cellular microenvironment is one of the key regulators of functions of SCs and their differentiation fate [76,77]. Unlike traditional dish-based culture techniques, microfluidic cultures of SCs offer the advantage of manipulating physical and chemical environment of SCs in a spatio-temporally controlled manner and setting up scalable and parallel experiments. Growth and differentiation of human neural stem cells (hNSCs) was explored in a gradient-generating microfluidic platform by Chung et al. [78]. Diffusive mixing in the laminar flow regime throughout a network of microchannels generated a gradient of a mixture of several growth factors (GFs). A cell culture area at the bottom of the device was exposed to this gradient. Continuous perfusion ensured autocrine and paracrine signaling was minimized. It was demonstrated that cell proliferation was proportional to the concentration of GFs, whereas differentiation to astrocytes was inversely proportional to GFs concentration. Preserving autocrine and paracrine factors is also possible if cell-cell communication via soluble factors is concerned [79]. Kim et al. described a microfluidic device for perfused culture of stem cells with logarithmic range of flow rates. Syringe-driven flow combined with a network of fluidic resistances generated logarithmic flow conditions through separate cell-culture chambers in a single device [80]. Murine embryonic stem cells (mESCs) were cultured and perfused with media at rates of 0.001 to 1.1 µl/min for four days. Morphology of mESCs was found to be strongly influenced by the flow rate. Cells at the slowest perfusion did not proliferate. By increasing the flow rate, mESCs formed larger but fewer numbers of colonies. Larger colony areas at higher flow rates were ascribed to increased nutrient delivery, increased waste removal, and increased removal of proliferation-inhibiting secreted factors. The device was further modified to accommodate both logarithmic perfusion rates and concentration gradients in one microfluidic chip, making it possible to simultaneously explore a wider range of biological activities.


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Figallo et al. described a micro-bioreactor comprising arrays of wells for the culture of different cell types including hESCs [81]. Each well was independently perfused with culture medium through a network of microchannels. To accommodate the culture of hESCs in a 3D setting, the wells were coated with a thin layer of a photopolymerizing hydrogel. The device allowed for controlled growth of hESCs under different flow configurations and cell densities and facilitated their differentiation into vascular cells. Recently, Zhong et al. developed a microfluidic platform to extract total mRNA from a single hESC and synthesize cDNA on the same device with a mRNA-to-cDNA efficiency superior to that of the bench-top counterparts [82]. The device may be useful for understanding the normal and pathological development of human cells and tissues. The growing interest in the microfluidic-based stem cell research over the past decade has led to the development of platforms with high levels of control over soluble microenvironment of SCs. Integration of heterotypic cultures and 3D cues is expected to thrust this area of research further. 6.2.2.4 Braille display-based microfluidic cell culture platforms

Transport of fluids relies on various pumping and valving mechanisms such as syringe pumps [83], gravity-driven pumps [55], and hydrogel valves [84]. Unfortunately, it is difficult to integrate such components into microfluidic devices. Furthermore, due to the slow nature of mixing in the laminar flow regime, these mechanisms cannot provide mixing and delivery of different reagents at various combinations and concentrations that is required for certain in vitro cell studies. A microfluidic plaform that utilizes programmed movement of arrays of pins of a Braille display (BD) as integrated valves and pumps was developed by Gu et al. [85]. Channel replicas are sealed against a thin PDMS sheet of ~140 µm. The network of microchannels is aligned onto a grid of Braille pin actuators. Valving action is generated by movement of a Braille pin up and down. Upward movement of a pin deforms the PDMS sheet and constricts the channel section immediately above it, whereas retracting the pin opens the constricted channel. Synchronous movement of three pins (valves) results in a peristaltic pump that moves fluids in a pulsatile fashion. This computerized fluidic system enables rapid mixing of several streams, multiple laminar flows with minimal mixing between them, and segmented flow of immiscible fluids (Fig. 6.5).

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Futai et al. improved the BD-based microfluidics by reformulating the culture media to eliminate the need for exogeneous CO2, incorporating a transparent heater in the device to locally heat the cell cultivation area, and stabilizing the device using a set of a monolithic aluminum plates and hold-down clamps [86]. Incubator-free culture of C2C12 myoblasts and MC3T3-E1 osteoblasts was demonstrated over a period of two weeks without medium exchange. The BD-based cell culture platform is suited for on-site cell culture applications due to its compactness, portablity, optically accessibility of the cell cultivation area, and control over fluidic transport.

Fig. 6.5 (Left) The configuration of Braille display-based microfluidic setup and (Right) Segmented flow of perfluorodecalin and red and green food dyes (Images reproduced, with permission, from ref. [85] Copyright (2004) National Academy of Sciences, U.S.A.)

6.2.2.5 Endothelium on a chip

Endothelial cells (ECs) line the inner surface of blood vessels and form an interface between blood flow in the lumen and the rest of the vessel wall. In vivo, ECs are subjected to hemodynamic shear stress of 5-20 dyn/cm2, which induces various cellular responses [87-89] including in the morphology of ECs such that cells become elongated and aligned in the direction of the blood flow [90,91]. Understanding the effect of mechanical forces such as shear stress on ECs is fundamental to the study of certain diseases such as thrombosis and atherosclerosis. A BD-based microfluidic endothelium culture platform was described by Song et al. to systematically study the influence of physiological levels of


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shear stress on ECs morphology [15]. Microchannels with different contact areas were incorporated to allow for the displacement of different volumes of media at a given pumping frequency (shown as small pump and large pump in Fig. 6.6), generating various levels of shear stress in a single device. A unique feature of this system is the generation of pulsatile fluid flow that represents the nature of blood flow in the arterial vasculature. ECs are known to discriminate between pulsatile and non-pulsatile flows [91], and hence in vitro EC culture systems must generate pulsatile flows to find physiologic value. ECs response to shear stress was evaluated using the angle of orientation (AO) (the angle between cell’s major axis and the flow direction) and the shape index (SI) (dimensionless number that is 1 for a straight line and 0 for a circle). For shear stresses of

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