Microchip Capillary Electrophoresis: Methods and Protocols [1 ed.] 1588292932, 1597450766, 9781588292933

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METHODS IN MOLECULAR BIOLOGY ™

339

Microchip Capillary Electrophoresis Methods and Protocols Edited by

Charles S. Henry

Microchip Capillary Electrophoresis

M E T H O D S I N M O L E C U L A R B I O L O G Y™

John M. Walker, SERIES EDITOR 352. Protein Engineering Protocols, edited by Kristian 352 Müller and Katja Arndt, 2006 351. C. elegans: Methods and Applications, edited by 351 Kevin Strange, 2006 350. Protein Folding Protocols, edited by Yawen Bai 350 and Ruth Nussinov 2006 349. YAC Protocols, Second Edition, edited by Alasdair 349 MacKenzie, 2006 348. Nuclear Transfer Protocols: Cell Reprogramming 348 and Transgenesis, edited by Paul J. Verma and Alan Trounson, 2006 347. Glycobiology Protocols, edited by Inka 347 Brockhausen-Schutzbach, 2006 346. Dictyostelium discoideum Protocols, edited by 346 Ludwig Eichinger and Francisco Rivero-Crespo, 2006 345. Diagnostic Bacteriology Protocols, Second Edition, 345 edited by Louise O'Connor, 2006 344. Agrobacterium Protocols, Second Edition: 344 Volume 2, edited by Kan Wang, 2006 343. 343 Agrobacterium Protocols, Second Edition: Volume 1, edited by Kan Wang, 2006 342. 342 MicroRNA Protocols, edited by Shao-Yao Ying, 2006 341. 341 Cell–Cell Interactions: Methods and Protocols, edited by Sean P. Colgan, 2006 340. 340 Protein Design: Methods and Applications, edited by Raphael Guerois and Manuela López de la Paz, 2006 339. 339 Microchip Capillary Electrophoresis: Methods and Protocols, edited by Charles S. Henry, 2006 338. 338 Gene Mapping, Discovery, and Expression: Methods and Protocols, edited by M. Bina, 2006 337. 337 Ion Channels: Methods and Protocols, edited by James D. Stockand and Mark S. Shapiro, 2006 336. 336 Clinical Applications of PCR, Second Edition, edited by Y. M. Dennis Lo, Rossa W. K. Chiu, and K. C. Allen Chan, 2006 335. 335 Fluorescent Energy Transfer Nucleic Acid Probes: Designs and Protocols, edited by Vladimir V. Didenko, 2006 334. 334 PRINS and In Situ PCR Protocols, Second Edition, edited by Franck Pellestor, 2006 333. 333 Transplantation Immunology: Methods and Protocols, edited by Philip Hornick and Marlene Rose, 2006 332. 332 Transmembrane Signaling Protocols, Second Edition, edited by Hydar Ali and Bodduluri Haribabu, 2006 331. 331 Human Embryonic Stem Cell Protocols, edited by Kursad Turksen, 2006 330. 330 Embryonic Stem Cell Protocols, Second Edition, Vol. II: Differentiation Models, edited by Kursad Turksen, 2006 329. 329 Embryonic Stem Cell Protocols, Second Edition, Vol. I: Isolation and Characterization, edited by Kursad Turksen, 2006

328. 328 New and Emerging Proteomic Techniques, edited by Dobrin Nedelkov and Randall W. Nelson, 2006 327 Epidermal Growth Factor: Methods and Protocols, 327. edited by Tarun B. Patel and Paul J. Bertics, 2006 326 In Situ Hybridization Protocols, Third Edition, 326. edited by Ian A. Darby and Tim D. Hewitson, 2006 325 Nuclear Reprogramming: Methods and Protocols, 325. edited by Steve Pells, 2006 324 Hormone Assays in Biological Fluids, edited by 324. Michael J. Wheeler and J. S. Morley Hutchinson, 2006 323 Arabidopsis Protocols, Second Edition, edited by 323. Julio Salinas and Jose J. Sanchez-Serrano, 2006 322 Xenopus Protocols: Cell Biology and Signal 322. Transduction, edited by X. Johné Liu, 2006 321. Microfluidic Techniques: Reviews and Protocols, 321 edited by Shelley D. Minteer, 2006 320. Cytochrome P450 Protocols, Second Edition, edited 320 by Ian R. Phillips and Elizabeth A. Shephard, 2006 319. 319 Cell Imaging Techniques: Methods and Protocols, edited by Douglas J. Taatjes and Brooke T. Mossman, 2006 318 Plant Cell Culture Protocols, Second Edition, edited 318. by Victor M. Loyola-Vargas and Felipe Vázquez-Flota, 2005 317 317. Differential Display Methods and Protocols, Second Edition, edited by Peng Liang, Jonathan Meade, and Arthur B. Pardee, 2005 316. Bioinformatics and Drug Discovery, edited by 316 Richard S. Larson, 2005 315. Mast Cells: Methods and Protocols, edited by Guha 315 Krishnaswamy and David S. Chi, 2005 314 314. DNA Repair Protocols: Mammalian Systems, Second Edition, edited by Daryl S. Henderson, 2006 313. Yeast Protocols, Second Edition, edited by Wei 313 Xiao, 2005 312. Calcium Signaling Protocols, Second Edition, 312 edited by David G. Lambert, 2005 311. Pharmacogenomics: Methods and Protocols, 311 edited by Federico Innocenti, 2005 310. 310 Chemical Genomics: Reviews and Protocols, edited by Edward D. Zanders, 2005 309. 309 RNA Silencing: Methods and Protocols, edited by Gordon Carmichael, 2005 308. 308 Therapeutic Proteins: Methods and Protocols, edited by C. Mark Smales and David C. James, 2005 307. 307 Phosphodiesterase Methods and Protocols, edited by Claire Lugnier, 2005 306. 306 Receptor Binding Techniques, Second Edition, edited by Anthony P. Davenport, 2005 305. 305 Protein–Ligand Interactions: Methods and Applications, edited by G. Ulrich Nienhaus, 2005

M E T H O D S I N M O L E C U L A R B I O L O G Y™

Microchip Capillary Electrophoresis Methods and Protocols

Edited by

Charles S. Henry Colorado State University, Fort Collins, CO

© 2006 Humana Press Inc. 999 Riverview Drive, Suite 208 Totowa, New Jersey 07512 www.humanapress.com All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise without written permission from the Publisher. Methods in Molecular BiologyTM is a trademark of The Humana Press Inc. All papers, comments, opinions, conclusions, or recommendations are those of the author(s), and do not necessarily reflect the views of the publisher. This publication is printed on acid-free paper. ∞ ANSI Z39.48-1984 (American Standards Institute) Permanence of Paper for Printed Library Materials. Cover illustration:Figure 1 from Chapter 4, "Fabrication of Polymer Microfluidic Systems by Hot Embossing and Laser Ablation," by Laurie E. Locascio, David J. Ross, Peter B. Howell, and Michael Gaitan. Production Editor: Erika J. Wasenda Cover design by Patricia F. Cleary For additional copies, pricing for bulk purchases, and/or information about other Humana titles, contact Humana at the above address or at any of the following numbers: Tel.: 973-256-1699; Fax: 973-256-8341; E-mail: [email protected]; or visit our website at www.humanapress.com. Photocopy Authorization Policy: Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Humana Press Inc., provided that the base fee of US $30.00 per copy is paid directly to the Copyright Clearance Center at 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license from the CCC, a separate system of payment has been arranged and is acceptable to Humana Press Inc. The fee code for users of the Transactional Reporting Service is: [1-58829-293-2/06 $30.00 ]. Printed in the United States of America. 10 9 8 7 6 5 4 3 2 1 E-ISBN 1-59745-076-6 ISSN 1064-3745 Library of Congress Cataloging-in-Publication Data Microchip capillary electrophoresis : methods and protocols / edited by Charles S. Henry. p. ; cm. -- (Methods in molecular biology ; 339) Includes bibliographical references and index. ISBN 1-58829-293-2 (alk. paper) 1. Capillary electrophoresis. 2. Integrated circuits. I. Henry, Charles S. (Charles Sherman), 1972- . II. Series: Methods in molecular biology ; v. 339. [DNLM: 1. Electrophoresis, Microchip. W1 ME9196J v.339 2006 / QU 25 M626 2006] QP519.9.C36M53 2006 572.8028--dc22 2005028755

Preface Microchip capillary electrophoresis emerged as an important new analytical technique in the early 1990s out of the pioneering work of a group of dedicated scientists at Ciba Geigy. This new technique was a result of the marriage of the ability of conventional capillary electrophoresis to analyze ultrasmall volumes (nL) and microfabrication techniques perfected in the semiconductor industry to produce very small structures in silicon. The resulting technology holds significant promise to improve our ability for analysis of biological systems because it can handle objects as small or smaller than a single cell, integrate such sample processing steps as filtration and the polymerase chain reaction, and generate answers in seconds or minutes compared with the hours used for many traditional techniques. The goal of this volume of Humana’s series Methods in Molecular BiologyTM is to provide the reader an overview of the methods currently in place for microchip capillary electrophoresis, as well as to provide useful practical information on how to get started in the field. The text of Microchip Capillary Electrophoresis is divided into four sections. Part I deals with fabrication methods for the production of microchips because this is fundamental to the ability to use the technology. The chapters are divided based on the substrate material and include glass (Chapter 2), poly(dimethylsiloxane) (Chapter 3), and other polymers including polymethylmethacrylate (Chapter 4). The information provided in these chapters should be suitable for even the novice to produce simple microchips for standard separations. Part II discusses methods to control the surface chemistry and measure the resulting alterations in microfluidic devices. Surface chemistry plays an important role in systems at this scale and must be carefully considered for optimal operation. Chapter 5 provides a general overview of several methods of both adsorbed and covalent surface modification. Chapter 6 provides more detail on a simple, yet effective, adsorbed coating system. Part III describes different detection modes for microchip capillary electrophoresis with detail provided for mass spectrometry (Chapter 7), electrochemistry (Chapter 8), and finally conductivity (Chapter 9). The last section of this book outlines applications of microchip capillary electrophoresis for biological analysis. Chapters 10–12 deal with the analysis of DNA, proteins, and peptides, respectively. Chapter 13 discusses techniques for measuring the impact of surface modification on flow in microfluidic channels. Chapter 14 discusses single cell analysis. The last chapter (Chapter 15) is a forward-looking review of the integration of the

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polymerase chain reaction into a capillary electrophoresis microchip, part of the next generation of a technology that is focused on the eventual integration of all laboratory functions in a single device. Microchip Capillary Electrophoresis is intended to be both a practical guide for those interested in using this exciting new technology in their own research, as well as an important source of fundamental information detailing how the technique works at the molecular scale. As such, our book is not meant to be an all inclusive, exhaustive text on every aspect of microchip capillary electrophoresis. Readers are encouraged to use the book as the reference guide it was intended to be, and then move on to seek current literature in this rapidly evolving field for applications more specific to their work. Finally, the editor would like to thank Professor John Walker for his significant help in completing this work. Suggestions for content from Drs. Scott Martin and Steve Soper are also acknowledged. Finally, the continued support of the publisher throughout this lengthy process was also greatly appreciated.

Charles S. Henry

Contents Preface .............................................................................................................. v Contributors .....................................................................................................ix 1 Microchip Capillary Electrophoresis: An Introduction Charles S. Henry ................................................................................... 1

PART I MICROCHIP FABRICATION METHODS 2 Fabrication of a Glass Capillary Electrophoresis Microchip With Integrated Electrodes Mark M. Crain, Robert S. Keynton, Kevin M. Walsh, Thomas J. Roussel, Jr., Richard P. Baldwin, John F. Naber, and Douglas J. Jackson ................................................................... 13 3 Micro-Molding for Poly(dimethylsiloxane) Microchips Carlos D. García and Charles S. Henry .............................................. 27 4 Fabrication of Polymer Microfluidic Systems by Hot Embossing and Laser Ablation Laurie E. Locascio, David J. Ross, Peter B. Howell, and Michael Gaitan ........................................................................ 37

PART II SURFACE MODIFICATION METHODS 5 Surface Modification Methods for Enhanced Device Efficacy and Function Barbara J. Jones and Mark A. Hayes ................................................... 49 6 Polyelectrolyte Coatings for Microchip Capillary Electrophoresis Yan Liu and Charles S. Henry ............................................................. 57

PART III DETECTION METHODS FOR MICROCHIP CAPILLARY ELECTROPHORESIS 7 Interfacing Microchip Capillary Electrophoresis With Electrospray Ionization Mass Spectrometry Trust Razunguzwa and Aaron T. Timperman ..................................... 67 8 Interfacing Amperometric Detection With Microchip Capillary Electrophoresis R. Scott Martin .................................................................................... 85 9 Conductivity Detection on Microchips Roland Hergenröder and Benedikt Graß .......................................... 113

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PART IV APPLICATIONS

Contents OF

MICROCHIP CAPILLARY ELECTROPHORESIS

10 DNA Separations Andrea W. Chow .............................................................................. 11 Protein Separations Andrea W. Chow .............................................................................. 12 Microchip Capillary Electrophoresis: Application to Peptide Analysis Barbara A. Fogarty, Nathan A. Lacher, and Susan M. Lunte ............ 13 Measuring Electroosmotic Flow in Microchips and Capillaries S. Douglass Gilman and Peter J. Chapman ....................................... 14 Single Cell Analysis on Microfluidic Devices Christopher T. Culbertson ................................................................ 15 Rapid DNA Amplification in Glass Microdevices Christopher J. Easley, Lindsay A. Legendre, James P. Landers, and Jerome P. Ferrance ................................................................

129 145

159 187 203

217 Index ............................................................................................................ 233

Contributors RICHARD P. BALDWIN • Department of Chemistry, University of Louisville, Louisville, KY PETER J. CHAPMAN • Department of Chemistry, University of Tennessee, Knoxville, TN ANDREA W. CHOW • Microfluidics Research and Development, Caliper Life Sciences, Mountain View, CA MARK M. CRAIN • Lutz Micro/Nanotechnology Cleanroom, University of Louisville, Louisville, KY CHRISTOPHER T. CULBERTSON • Department of Chemistry, Kansas State University, Manhattan, KS CHRISTOPHER J. EASLEY • Department of Chemistry, University of Virginia, Charlottesville, VA JEROME P. FERRANCE • Department of Chemistry, University of Virginia, Charlottesville, VA BARBARA A. FOGARTY • Life Sciences Interface, Tyndall Institute, Cork, Ireland MICHAEL GAITAN • Analytical Chemistry Division, National Institute of Standards and Technology, Gaithersburg, MD CARLOS D. GARCÍA • Department of Chemistry, University of Texas– San Antonio, San Antonio, TX S. DOUGLASS GILMAN • Department of Chemistry, Louisiana State University, Baton Rouge, LA BENEDIKT GRAß • ISAS-Institute for Analytical Sciences, Dortmund, Germany MARK A. HAYES • Department of Chemistry, Arizona State University, Tempe, AZ CHARLES S. HENRY • Department of Chemistry, Colorado State University, Fort Collins, CO ROLAND HERGENRÖDER • ISAS-Institute for Analytical Sciences, Dortmund, Germany PETER B. HOWELL • Analytical Chemistry Division, National Institute of Standards and Technology, Gaithersburg, MD DOUGLAS J. JACKSON • Department of Electrical and Computer Engineering, University of Louisville, Louisville, KY BARBARA J. JONES • Analytical Chemistry Division, National Institute of Standards and Technology, Gaithersburg, MD

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ROBERT S. KEYNTON • Department of Bioengineering, University of Louisville, Louisville, KY NATHAN A. LACHER • Analytical Research and Development, Pfizer Global Biologics, St. Louis, MO JAMES P. LANDERS • Department of Chemistry, University of Virginia, Charlottesville, VA LINDSAY A. LEGENDRE • Department of Chemistry, University of Virginia, Charlottesville, VA YAN LIU • Department of Chemistry, Colorado State University, Fort Collins, CO LAURIE E. LOCASCIO • Analytical Chemistry Division, National Institute of Standards and Technology, Gaithersburg, MD SUSAN M. LUNTE • Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, KS R. SCOTT MARTIN • Department of Chemistry, Saint Louis University, St. Louis, MO JOHN F. NABER • Department of Electrical and Computer Engineering, University of Louisville, Louisville, KY TRUST RAZUNGUZWA • Department of Chemistry, West Virginia University, Morgantown, WV DAVID J. ROSS • Analytical Chemistry Division, National Institute of Standards and Technology, Gaithersburg, MD THOMAS J. ROUSSEL, JR. • Department of Bioengineering, University of Louisville, Louisville, KY AARON T. TIMPERMAN • Department of Chemistry, West Virginia University, Morgantown, WV KEVIN M. WALSH • Department of Electrical and Computer Engineering, University of Louisville, Louisville, KY

1 Microchip Capillary Electrophoresis An Introduction Charles S. Henry Summary Microchip capillary electrophoresis emerged in the early 1990s as an intresting and novel approach to the high-speed separation of biological compounds, including DNA and proteins. Since the early development in this area, growth in the research field has exploded and now includes chemists and engineers focused on developing new and better microchips, as well as biologists and biochemists who have begun to apply this exciting and still relatively new methodology to real-world problems. This chapter seeks to outline the historical development of microchip, the key elements of microchip capillary electrophoresis, and finally some of the important applications beign develop that utilize microchip capillary electrophoresis. Key Words: Microchip capillary electrophoresis; capillary electrophoresis; microfabrication; bioanalytical chemistry.

1. Introduction Microchip capillary electrophoresis (CE) has appeared over the decades as a result of the marriage of chemical analysis and microfabrication techniques from the integrated circuit world. The concept of microchip separations is not new, however, with the first report of a microfabricated gas chromatography column appearing in 1979 (1) and the first example of a microfabricated liquid chromatography system appearing in 1990 (2). Little excitement was generated over these early developments, however, because of the complexity of the operational systems and generally poor performance of the devices. Active development of microchip separation technology did not begin until the early 1990s with the seminal work of Manz, Harrison, Verpoorte, and Widmer in microchip CE (3). CE proved to be an excellent match for microchip technologies because it easily manipulates volumes at the nanoliter scale, requires no moving parts, and provides fast, high-resolution separations. From: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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Since the early reports of microchip CE, the field has grown exponentially in both the number of investigators and the areas of application and fundamental development. One general theme now driving the field is the integration of function, with the ultimate goal being the development of miniaturized total analysis systems (also referred to as Lab-on-a-Chip) that integrate all functions of the modern analytical laboratory in a single device. Unlike traditional CE instrumentation, which consists of essentially a single capillary (or an array of capillaries in parallel), many different capillaries and fluidic channels can be patterned on a microfluidic device, providing the potential for high-throughput, massively parallel analysis. Microchip CE has also garnered significant attention because of the potential applications and overall device performance. Although the primary early focus in the field was on DNA analysis (4), microchip CE has been rapidly adapted to many biological, environmental, and industrial applications (5–12). Microchip CE also has the added benefits of low cost, small size, and fast analysis times, which are general goals of many chemical analysis methods. Functionality, such as polymerase chain reaction (PCR), enzymatic digestion, and solid phase extraction, can be incorporated into the microchip to provide pre- and postsample processing. This capability is important because it allows a raw sample to be added to the microchip and a final quantitative answer provided. 2. Theory and Mechanisms of Action in Microchip CE The theory and mechanisms of action for microchip CE are based on fundamental knowledge gained in the development of conventional CE. The first report of conventional CE is generally attributed to Lukacs and Jorgenson in the early 1980s (13). In this work, mixtures of proteins and peptides were separated in glass capillaries. The two major outcomes of this technology were (1) the demonstration of open tubular electrophoresis for high-speed separations of biomolecules and (2) the presence of electroosmotic flow (EOF). EOF is a bulk solution flow phenomenon that occurs in capillaries filled with mild ionic solutions (typically 100) and resolution are needed, the use of other radiation sources like X-rays, electron beams, or excimer lasers is recommended.

References 1. Ng, J. M., Gitlin, I., Stroock, A. D., and Whitesides, G. M. (2002) Components for integrated poly(dimethylsiloxane) microfluidic systems. Electrophoresis 23, 3461–3473. 2. Becker, H. and Locascio, L. (2002) Polymer microfluidic devices. Talanta 56, 267–287. 3. Duffy, D. C., McDonald, J. C., Schueller, O. J. A., and Whitesides, G. M. (1998) Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70, 4974–4984. 4. Lin, C. -H., Lee, G. -B., Chang, B. -W., and Chang, G. -L. (2002) A new fabrication process for ultra-thick microfluidic microstructures utilizing SU-8 photoresist. J. Micromech. Microeng. 12, 590–597. 5. Unger, M. A., Chou, H. -P., Thorsen, T., Scherer, A., and Quake, S. R. (2000) Monolithic microfabricated valves and pumps by multilayer soft lithography. Science 288, 113–116. 6. Khoury, C., Mensing, G. A., and Beebe, D. J. (2002) Ultra rapid prototyping of microfluidic systems using liquid phase photopolymerization. Lab on a Chip 2, 50–55. 7. Chrisey, D. B. and Pique, A. (2002). Introduction to Direct-Write technologies for rapid prototyping, in Direct-Write Technologies for Rapid Prototyping Applications: Sensors, Electronics, and Integrated Power Sources (Pique, A. and Chrisey, D. B., eds.), Academic Press, San Diego, CA, pp. 1–16.

4 Fabrication of Polymer Microfluidic Systems by Hot Embossing and Laser Ablation Laurie E. Locascio, David J. Ross, Peter B. Howell, and Michael Gaitan Summary Fabrication of microfluidic channels in common commercially available thermoplastic materials can be easily accomplished using hot embossing or ultraviolet (UV) laser ablation. Hot embossing involves replication of a microfluidic network in a polymer substrate from a stamp (or template) fabricated in silicon or metal. UV laser ablation is performed by either exposing the polymer substrate through a mask or by using a laser direct-write process. The resulting polymer microfluidic channels are most often sealed with another polymer piece using thermal bonding or solvent bonding to complete the fabrication procedure. Unlike their silicon and glass counterparts, polymer microfluidic systems can be fabricated by these methods in less than 1 h, making the materials attractive for both research prototyping and commercialization. Key Words: Hot embossing; laser ablation; polymer; microfluidics; thermoplastic; bonding; excimer laser.

1. Introduction In the late 1990s, several new techniques were introduced for the fabrication of polymer microchannels including hot embossing (1), laser ablation (2), X-ray photolithography (3), injection molding (4), and soft embossing (5). Polymers have distinct advantages over other substrates for many applications because of their low cost and the ease with which they can be micromachined. In fact, techniques such as polymer embossing and laser ablation made microfluidics technology widely accessible, thus, greatly expanding the field of microfluidics in a short time. Our group was one of the first to introduce methods for hot embossing and room temperature embossing (stamping, imprinting), and we have demonstrated their usefulness for fabricating microchannel networks in a variety of thermoplastic materials (1). During the hot embossing process, a polymer is pressed against a stamp (template) while heating to a temperature greater than From: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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the glass transition temperature allowing the polymer to flow and, thus, leaving an imprint of the stamp feature. In the room temperature embossing process, a polymer is pressed against a stamp under higher pressure, thus transferring the feature without allowing the polymer to flow and then reform. Both methods of embossing permit rapid prototyping of microchannels that can be accomplished in minutes. The limiting step in this process is the fabrication of the micromachined stamp or template. Once fabricated, however, the stamp can be used repeatedly to emboss hundreds of polymer microchannel networks. For rapid device prototyping, laser ablation (2) is an attractive alternative to embossing because it does not require the fabrication of an embossing template. Potential drawbacks to laser ablation are that the ablated channel surfaces are typically much rougher than embossed surfaces, and the surface chemistry (hydrophobicity, surface charge) of ablated channels is very different from that of the native sheet plastic. The modification of surface chemistry by laser ablation can be used to advantage, however, as it allows for the potential machining and chemical modification of microfluidic channels in one step. The surface chemistry resulting from ablation can even be manipulated through variation of the atmosphere surrounding the plastic during ablation (6). 2. Materials (see Note 1) 1. For hot embossing, commercial thermoplastic sheet such as polycarbonate (PC), polystyrene (PS), or polymethylmethacrylate (PMMA) is used. Also required for this procedure are a micromachined template or stamp, a hydraulic press, and two polished temperature-controlled heating blocks. The materials and methods are described in detail next for the silicon micromachining; however, this requires a clean room and microfabrication capabilities. Alternatively, the template can be fabricated at fully equipped microfabrication facilities that provide contract services. 2. Materials required for laser micromachining include polymer or glass substrates (there is much more flexibility in the choice of material when using laser ablation). Also needed are a laser ablation system with a pulsed UV excimer laser, a motorized translation stage, optics to shape and direct the laser output to the polymer surface, and a computer to control laser firing and stage movement. 3. For polymer channel sealing, materials include a drill, microscope slides, binder clips, and a circulating air oven.

3. Methods 3.1. Polymer Microchannel Fabrication by Hot Embossing Microfluidic channels are fabricated by hot embossing using a micromachined silicon wafer (1) as a stamp or template. The fabrication procedure for the micromachined silicon template is described in detail in Subheading 3.1.1., followed by the procedure for making polymer microfluidic systems from these templates.

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3.1.1. Silicon Template Micromachining 1. Low-doped (100) silicon wafers are placed in a wet oxidation furnace at 980°C for 20 min to create a 100-nm thick oxide film. 2. Photoresist is spun-on the wafer and the wafer is soft baked. Hexamethyldisilazane is spun-on before application of the photoresist as an adhesion promoter. In addition, photoresist is spun-on to the backside of the wafer. Finally, the wafers are rinsed in deionized (DI) water and spin dried. 3. A photomask is prepared for patterning the photoresist. For this work, our designs generally require a 20-µm minimum feature size. We create the photomasks by high-resolution printout onto clear transparency film. Alternatively, chromium photomasks are fabricated for us at the University of California, Berkeley Microfabrication Laboratory (http://microlab.berkeley.edu/) when smaller features are required. The mask contains the drawings of the fluid channel elements and an alignment mark for rotational alignment to the flat of the silicon wafer. All features are drawn in Manhattan format, i.e., lines or boxes parallel or perpendicular to the alignment mark, so that they will be aligned to the (111) planes of the silicon wafer. This alignment is required for anisotropic etching of the silicon wafer. 4. The mask is placed over the front side of the silicon wafer and aligned to the flat using the alignment mark. The photoresist is exposed using an UV light source and patterned using a developer, then rinsed and dried. 5. Once the photoresist is patterned, it is used to pattern the silicon dioxide film by a 2 min wet etch in a 6% (volume fraction) buffered hydrogen fluoride (HF) oxide etch solution, then rinsed and dried. Following this, the photoresist is stripped in acetone, and then the wafer is again rinsed and dried. 6. The silicon wafers are then anisotropically etched in tetramethylammonium hydroxide (TMAH) solution. We prepare a TMAH etch solution, based on the work reported by Tabata et al. (7), consisting of 450 mL of 25% by weight electronic grade TMAH in aqueous solution mixed with 900 mL of DI water (one part 25% TMAH to two parts DI water). The solution is placed in a reflux etch container and heated to 80°C on a hotplate. The wafers are placed in a Teflon holder and immersed in the solution. Following the work by Klaassen et al. (8), 1 g of ammonium peroxydisulfate (APODS) in powder form is added every 10 min during etching in order to eliminate the formation of hillocks. The etch rate for this solution in the (100) direction was measured as 0.9 µm/min. 7. Finally, the silicon dioxide mask is stripped by a second 2 min wet etch in 6% buffered HF, then rinsed and dried. The cross-section of the raised features on the template are trapezoidal-shaped as depicted in Fig. 1, with a width at the top surface ranging between 20 and 100 µm depending on our application. Smaller template features can be made using the Cr masks. The sides of the trapezoid are aligned to the (111) planes of the silicon crystal forming a 54.74° angle from the wafer surface plane (9). 8. The silicon template may be used to emboss channels, or the micromachined silicon template may be used as a master to fabricate a more robust template in metal (4). In the second case, a metal electroform is made from the silicon master that is

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Fig. 1. Micromachined silicon template for channel embossing. the mirror image of the silicon template. Then, a second metal electroform is created from the first electroform that is an exact replica of the original silicon template. Thus, micrometer features are transferred to a more robust metal substrate that can be used to fabricate microchannels in plastic substrates by hot embossing or injection molding.

3.1.2. Polymer Embossing We have used the etched silicon template to emboss microchannels in plastic materials at room temperature (10), or at elevated temperatures (hot embossing) (11). The hot embossing procedure is as follows: 1. Many types of commercially available sheet plastic, such as PS, PC, PMMA, polyethylenetetraphthalate glycol (PETG), and polyvinylchloride (PVC), have been used to fabricate polymer microchannels (1,10,12–15). In our lab, the sheet plastic is cut into rectangular pieces that are approx 2.0 × 6.0 cm. A cut plastic piece is washed with ethanol and then dried under nitrogen, and placed on top of a clean silicon template in a laminar flow hood. 2. The silicon template and plastic piece are placed between two polished aluminum blocks, one on the top and one on the bottom, each with its own embedded heater connected to a common temperature controller. The aluminum blocks are heated to a temperature either equal to or slightly greater than the glass transition temperature of the plastic material, but below the melt temperature. 3. The assembly composed of the aluminum blocks sandwiching the silicon template and plastic piece is placed in a hydraulic press. Pressure (2.8 × 106 Pa to 2.1 × 107 Pa, 400–3000 psi) is applied for a time that ranges typically from 5 to 60 min depending

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Table 1 Hot Embossing Parameters for Various Polymer Substrates Polymer substrate PMMA PETG PC

Temperature (°C)

Time (min)

Pressure (Pa)

110 80 155

60 20 90

5.1 × 106 (740 psi) 3.4 × 106 (500 psi) 1.4 × 107 (2000 psi)

Fig. 2. Hot embossed microfluidic channel in polymethylmethacrylate. on the plastic used and the embossing temperature. After the required time, the pressure is released, and the top aluminum block is removed. Typical conditions for imprinting polymer microchannels are described in Table 1. 4. The silicon template and the plastic piece are removed together with tweezers from the bottom aluminum block while at elevated temperature, and are placed on the bench top for cooling. Upon cooling, the plastic piece releases rapidly from the silicon template owing to a mismatch in the thermal expansion coefficients of the two materials. The resulting plastic piece contains microchannels that are the exact mirror of the raised features on the silicon template as shown in Fig. 2. 5. If the plastic piece warps significantly upon rapid cooling and release, the assembly (aluminum blocks, silicon template, plastic piece) can be cooled to approx 10°C below the glass transition temperature while still under pressure. Then pressure is released and the top aluminum block is removed. The silicon template and the plastic piece are removed from the bottom block and cooled to room temperature on the bench top as stated previously.

Alternatively, plastic devices can be imprinted at room temperature, rather than at elevated temperatures, and at higher pressures. When devices are imprinted at room temperature, the microchannel depth is dependent on imprinting pressure,

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imprinting time, and properties of the plastic; therefore, the resulting channel may not be an exact inverted replica of the silicon template. An advantage of room temperature imprinting, however, is that fabrication time is significantly reduced as compared with hot embossing. Reproducible imprints can be made at room temperature in less than 2 min (9). 3.2. Polymer Microchannel Fabrication by Laser Ablation A typical direct-write laser ablation system similar to the one in our laboratory consists of a pulsed UV excimer laser, a motorized translation stage that holds the substrate to be ablated, the optics to shape and direct the laser output to the surface to be ablated, and a computer to coordinate and control the translation stage and the firing of the laser. During ablation, the laser is fired with a repetition rate of 100 Hz, and the translation stage is used to move the plastic substrate beneath the laser to form microchannels. The basic procedure for direct-write laser ablation of a microfluidic device using a system from Potomac Photonics (see Note 1) (Lanham, MD) is as follows: 1. The laser system is first turned on and warmed up, and the laser cavity gas is changed, if necessary. Changing the gas and firing the laser for a few minutes at 100 Hz helps to ensure a more constant laser power and hence more uniform channel depths. We use either ArF or KrF gas to ablate at 193 or 248 nm, respectively. 2. The desired beam-defining aperture is installed, and the laser power is adjusted for the desired depth of cut. The aperture, along with the system optics, is used to set the shape and width of the ablated channels. The depth of the channels is determined by the material, laser pulse power (and wavelength), the repetition rate, and the feed rate of the translation stage. In most of our work, 50–60 µm wide by 50–100 µm deep channels are cut in commercially available sheet plastics using a round aperture, a power of 65 µJ/pulse, a repetition rate of 100–200 Hz, and a feed rate of 50 µm/s. Figure 3 shows examples of polymer microchannels ablated using these parameters. 3. The geometry of the microchannel network for the device to be fabricated is determined by a simple coded program that specifies sequences of translation stage movements and laser firings. New device designs can be created or old designs modified with as little as a few minutes of programming. 4. The plastic substrate is affixed to the translation stage, and the program is started to cut the device. Using the settings given in ref. 2, a device can be ablated in between 5 min and 2 h depending on the complexity. 5. The ablation process can create a great deal of particulate matter that can be redeposited onto the substrate and in the newly formed microchannels. After ablation, much of this debris is removed using a combination of rinsing and sonication in water, buffer, ethanol, or a mixture of the three. Once the particulate matter is removed, the device is ready for sealing using the same techniques applicable to embossed channels.

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Fig. 3. Microchannel cross-sections for laser ablation in various sheet polymers. All of the channels were cut using the same laser settings: a round aperture to define a 60-µm spot at the surface of the polymer, a laser power of 65 µJ/pulse, a laser repetition rate of 200 Hz, and a translation rate of 50 µm/s.

3.3. Sealing Polymer M icrochannels by Thermal Bonding

Polymer devices can be sealed by a number of techniques that include thermal bonding and solvent bonding. In our laboratories, most devices are sealed by thermal bonding processes that are described next. 1. A polymer piece that serves as the lid of the device is cut from sheet plastic to be approximately the same dimensions as the substrate containing the embossed or ablated channels. The polymer lid can be cut from the same substrate material that was used to form the microchannels. Alternatively, for easier sealing, the polymer lid can be cut from a different material with a lower glass transition temperature than that used to create the microchannels. 2. Holes are drilled into the lid to provide fluidic access. After drilling, the lid substrate is thoroughly rinsed in water, then ethanol, and dried under a compressed nitrogen jet to remove particulates. 3. The lid is placed over the microchannel substrate aligning the holes in the lid to the ends of the microchannels. The two substrates, lid and microchannel, are clamped together between four microscope glass slides, two on each side, using binder clips. The device is then thermally bonded in a circulating air oven. It is important to keep the time and temperature as low as possible during the sealing process to avoid physical alteration of the microchannel feature. Typical sealing times and temperatures for three common polymers are given in Table 2.

3.4. Polymer M icrochannel Fabrication by H ot Embossing 1. When hot embossing, the polymer substrate should fit entirely on the silicon substrate. If the polymer extends over the edge of the silicon template during heating,

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Sealing temperature (°C)

Time (min)

103 75 135

12 10 60

the polymer can deform around the edges of the silicon template, thus preventing its removal once the process is complete. 2. When hot embossing, pressure should not be applied in the hydraulic press until the desired temperature is reached. Applying pressure at cooler temperatures may result in silicon template fracture. Careful handling of the silicon template during hot embossing will permit repeated usage of the template for the fabrication of hundreds of plastic microfluidic devices. 3. When room temperature imprinting, the polymer piece should be larger than the silicon template. If the polymer substrate does not cover the template completely, the template will crack when pressure is applied. When embossing at room temperature, the lifetime of a silicon template is much shorter than with hot embossing and the template is subject to fractures, particularly with harder plastics such as PMMA and PC. Much longer template lifetimes can be achieved when embossing softer plastics, such as PVC or PETG.

3.5. Polymer Microchannel Fabrication by Laser Ablation 1. During laser ablation, it is important to keep the lens-to-polymer distance constant and to maintain constant laser power. 2. When ablating deep channels, the laser will defocus, thus changing the geometry of the ablated feature. 3. When ablating cross intersections, the point in the middle of the intersection will be approximately two times as deep as the surrounding microchannels unless great care is taken to prevent this by modifying laser parameters when ablating the intersection.

3.6. Sealing Polymer Microchannels by Thermal Bonding 1. When sealing polymer microchannels using thermal bonding, binder clips should be aligned directly over the channels so that air is not entrapped in the vicinity of the microchannel. 2. If the microchannel sealing is difficult to accomplish without melting the embossed feature, a polymer with a lower glass transition temperature can be used to seal the device. In this case, the sealing temperature is approximately the glass transition temperature of the lid material and not the channel material. 3. Cooling to room temperature while the template and plastic piece are under pressure will break the silicon template. Always release the pressure when cooling to a temperature that is less than 10°C below the polymer glass transition temperature.

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4. Good air circulation is required to maintain good temperature control during sealing. The best results are obtained in a circulating air oven such as a gas chromatography oven. 5. Room temperature imprinted channels cannot generally be sealed using thermal bonding. The polymer will often flow during the bonding process and destroy the feature during this type of sealing process. 6. After drilling the fluid access holes in the device lid, it is important to remove all particulates that are on the edge of the hole as a result of drilling. If the edge is not completely clear of particulates, or if the edge is rough, the end of the microchannel will become occluded during sealing.

4. Notes 1. Certain commercial equipment, instruments, or materials are identified in this report to adequately specify the experimental procedure. Such identification does not imply recommendation or endorsement by the National Institute of Standards and Technology, nor does it imply that the materials or equipment identified are necessarily the best available for the purpose.

References 1. Martynova, L., Locascio, L. E., Gaitan, M., Kramer, G. W., Christensen, R. G., and MacCrehan, W. A. (1997) Fabrication of plastic microfluid channels by imprinting methods. Anal. Chem. 69, 4783–4789. 2. Roberts, M. A., Rossier, J. S., Bercier, P., and Girault, H. (1997) UV laser machined polymer substrates for the development of microdiagnostic systems. Anal. Chem. 69, 2035–2042. 3. Soper, S. A., Ford, S. M., Qi, S., McCarley, R. L., Kelly, K., and Murphy, M. C. (2000) Polymeric microelectromechanical systems. Anal. Chem. 72, 642A–651A. 4. McCormick, R. M., Nelson, R. J., AlonsoAmigo, M. G., Benvegnu, J., and Hooper, H. H. (1997) Microchannel electrophoretic separations of DNA in injectionmolded plastic substrates. Anal. Chem. 69, 2626–2630. 5. Duffy, D. C., McDonald, J. C., Schueller, O. J. A., and Whitesides, G. M. (1998) Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70, 4974–4984. 6. Pugmire, D. L., Waddell, E. A., Haasch, R., Tarlov, M. J., and Locascio, L. E. (2002) Surface characterization of laser-ablated polymers used for microfluidics. Anal. Chem. 74, 871–878. 7. Tabata, O., Asahi, R., Funabashi, H., Shimaoka, K., and Sugiyama, S. (1992) Anisoptropic etching of silicon in TMAH solutions. Sens. Actuators A Phys. 34, 51–57. 8. Klassen, E. H., Reay, R. J., Storment, C., et al. (1996) Micromachined Thermally Isolated Circuits. Proc. Solid-State Sensor and Actuator Workshop, 127–131. 9. Seidel, H. (1987) The Mechanism of Anisotropic Silicon Etching and its Relevance for Micromachining. Proc. Transducers 87, 120–125. 10. Xu, J. D., Locascio, L., Gaitan, M., and Lee, C. S. (2000) Room-temperature imprinting method for plastic microchannel fabrication. Anal. Chem. 72, 1930–1933.

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11. Johnson, T. J., Waddell, E. A., Kramer, G. W., and Locascio, L. E. (2001) Chemical mapping of hot embossed and UV laser ablated microchannels in poly(methyl methacrylate) using carboxylate specific fluorescent probes. Appl. Surf. Sci. 181, 149–159. 12. Henry, A. C., Waddell, E. A., Shreiner, R., and Locascio, L. E. (2002) Control of electroosmotic flow in laser-ablated and chemically modified hot imprinted poly(ethylene terephthalate glycol) microchannels. Electrophoresis 23, 791–798. 13. Becker, H. and Heim, U. (2000) Hot embossing as a method for the fabrication of polymer high aspect ratio structures. Sens. Actuators A Phys. 83, 130–135. 14. Ueno, K., Kitagawa, F., Kim, H. B., et al. (2000) Fabrication and characteristic responses of integrated microelectrodes in polymer channel chip. Chem. Letters 8, 858, 859. 15. Uchiyama, K., Xu, W., Yamamoto, M., Shimosaka, T., and Hobo, T. (1999) Development of imprinted polymer microchannel capillary chip for capillary electrochromatography. Anal. Sciences 15, 825, 826.

II SURFACE MODIFICATION METHODS

5 Surface Modification Methods for Enhanced Device Efficacy and Function Barbara J. Jones and Mark A. Hayes Summary Currently available microfluidic devices can accomplish a variety of tasks useful in molecular biology. When moving analytical processes to a microenvironment, the properties of the device surface play a larger role in the functioning of the device. Surface modification may become necessary or advantageous for the purpose of control of the functional mechanics of the device, keeping cell components from adsorbing, attaching antibodies to the surface for detection of biological components, and attaching a functional bonding complex. Modification of the surface of microfluidic devices for the control of flow and device function, or for funtionalization of the surface to tailor the device to a specific use, can be accomplished in numerous bench-top, postfabrication procedures. The use of polyelectrolyte multilayers, ultraviolet grafting of polymers, and polydimethylsiloxane/surfactant coating to control flow and mitigate adsorption is discussed. In addition, the funtionalization of devices through amine termination of surfaces, and immobilization of biotin within a phosphotidylcholine bilayer is detailed. Key Words: Surface modification; biotinylation; microfluidics; polyelectrolyte multilayer.

1. Introduction Microfluidic devices will be used by every molecular biologist at some point in their career. Is that a wild and overstated claim? Probably not. There are a myriad of reasons why microfluidic devices will be commonplace. They reduce reagent needs, waste production, and time; the financial savings alone make the adaptation of devices to your particular needs worth exploring. The commercial devices already available can accomplish a variety of tasks, from protein separation and DNA sequencing, to combinatorial chemistry. One key aspect of microfluidic devices is that the samples are necessarily small and the ratio of the volume to the surface area shrinks significantly. As a consequence of this, the surface properties become important and certain enhancements may become necessary. Surface modification plays four overlapFrom: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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ping roles. Control of the surfaces offers the best means of control for the functional mechanics of the device. Modifications such as dynamic and covalently bound coatings will keep proteins and “gooey” cell components from sticking (1–7). Attachment of antibodies to the surface will allow you to better detect biological components (8,9). And finally, attaching a functional bonding complex to the surface makes individualized attachments possible (4). All of this can be done on your bench-top specific to your desired application. In addition to using small samples, microchips are fabricated in a wide array of materials, and specific surface modification may indeed be necessary for the optimization of your method. There are many modification methods, some of which are detailed here, that will help you to effect better separation of biological components and also provide the ability to modify your device to perform your specific analysis. The purposes for modifying the surface of microfluidic devices fall into two main categories: improvement of the surface properties for the normal function of the device, and funtionalization of the surface to tailor the device to a specific use. This chapter will provide several techniques for each. 1.1. Improvement of Surface Properties of Microfluidic Devices Improvement of surface properties may be necessary in any of the devices that you choose, whether you purchase commercially available devices or fabricate them yourself. Devices produced in polymers, such as poly(dimethylsiloxane) (PDMS) or polycarbonate, tend to be hydrophobic in character and, thus, loading of the channels can prove difficult. These polymer surfaces can also have poorly controlled electroosmotic flow (EOF; a property that greatly affects the separation efficiency and precision of a device; see Chapter 13). In addition, adsorption of biological components to the surface and even migration into the polymer matrix makes surface modification necessary. A promising technique that will be detailed here is the application of polyelectrolyte multilayers (PEMs) to these polymer device channel surfaces (2,7). This technique provides a simple and reproducible way to modify the surface of any polymer-based microfluidic device to provide a wettable channel with reproducible and stable EOF. An increased benefit to surface modification using this technique is the reproducibility of the EOF across devices fabricated from different polymers. The application of PEMs is easily achieved. The technique consists merely of filling the channel successively with alternating solutions of positive and negative polyelectrolytes allowing for the multilayers to form electrostatic bonds. Though these layers are not covalently bound to the surface, the multilayers provide complete coverage and remain robust even after long-term storage. A second technique for imposing a hydrophilic character on PDMS surfaces involves the ultraviolet (UV) grafting of polymers to the surface of the

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channels (5). This grafting is achieved by first creating grafting sites, radicals, at the surface by exposing the surface to UV irradiation while simultaneously exposing the device to a monomer solution. The monomers quickly react to form a polymer covalently bound at the reactive site. Devices fabricated in glass are highly charged on the surface and proteins often adsorb to these surfaces, making separations difficult. The manipulation of this surface charge and thus the EOF through changes in buffer pH can be useful; however, this variability in EOF may be a hindrance in many applications. To control EOF and decrease adsorption of proteins and other biological components, a PDMS/surfactant coating of glass microdevices can be utilized (1). 1.2. Funtionalization of Surfaces Funtionalization of the surfaces describes the immobilization of a component to the surface of a device, which then provides an available functional group for further bonding. This type of surface preparation is useful when developing assays or combinatorial reactions within the channels of the microdevice. There are many ways to functionalize devices; however, this discussion will be limited to funtionalization that is easily produced in postfabricated devices. The modification of poly(methylmethacrylate) (PMMA) devices to provide an amine-terminated surface has been demonstrated (4). These surfaces are then available for attachment of any variety of functional groups or targets. The procedure for amine termination can be accomplished on benchtop with little procedural complexity. The PMMA device pieces are first exposed to N-lithiodiaminopropane (N-LDAP). The reaction is then simply quenched with water. The amineterminated surface can additionally be exposed to octadecane chains to produce a highly ordered monolayer on the channel surfaces. Immobilization of biotin within a phosphotidylcholine bilayer on the surface of either PDMS or glass microdevices allows the streptavidin–biotin conjugation to be exploited in any desired utility. In this method, vesicles that contain a small percentage of biotinylated lipids are infused into the channels of either PDMS or glass microdevices where the vesicles adsorb to the surface and fuse to form a uniform bilayer coating (9). The biotin headgroups are then available at a number of surface sites. Any enzyme or reagent of interest that can be linked to streptavidin can thus be flowed over the biotinylated surface and immobilized at these sites. 2. Materials 2.1. Polyelectrolyte Multilayer (Option 1) 1. Dextran sulfate (MWav 5000, DS), 3% solution in ultrapurified water. 2. Hexadimethrine bromide (Polybrene, PB), 5% solution in ultrapurified water. 3. 0.1 M NaOH.

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2.2. Polyelectrolyte Multilayer (Option 2) 1. Polystyrene microdevice top section with imprinted channels and a PDMS cover film. 2. 60 mM poly(styrene sulfonate) sodium salt (PSS) (MWav 500,000) in 0.5 M NaCl adjusted to pH 9.0 with NaOH. 3. 20 mM poly(allylamine hydrochloride) (PAH) (MWav 70,000) in 0.5 M NaCl adjusted to pH 9.0 with NaOH. 4. 1 M NaOH. 5. All water used is ultrapurified (18 MΩ cm).

2.3. UV Graft Polymerization 1. Microfluidic device fabricated in PDMS. 2. Any of the following monomers: a. Acrylic acid (AA). b. Acrylamide (AM). c. Dimethylacrylamide (DMA). d. 2-hydroxylethyl acrylate (HEA). 3. Benzyl alcohol. 4. Sodium periodate (NaIO4).

2.4. PDMS Coating on Glass Microchips 1. 2. 3. 4. 5. 6.

Glass microchip with channels. 0.1 M NaOH. Dichloromethane (dried and distilled). PDMS 200. 0.01% Tween-20. Dry N2 gas.

2.5. Amine-Terminated PMMA 1. 2. 3. 4. 5. 6. 7.

PMMA microdevice, top and bottom separated. N-LDAP. n-Octadecane-1-isocynate (99%). Hexanes. Toluene. Acetone. Dry N2 gas.

2.6. Immobilization of Biotin Within a Phosphotidylcholine Bilayer 1. PDMS microdevice. 2. 1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC) (Avanti Polar Lipids, Inc., Alabaster, AL). 3. N-Biotinyl-CAP-PE (Avanti Polar Lipids). 4. Chloroform. 5. Phosphate buffer (pH 7.4, ionic strength 150 mM).

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6. 5 mg/mL bovine serum albumin (BSA) in pH 3.8 phosphate buffer solution. 7. NaCO3/NaHCO3 buffer (pH 9.8, ionic strength 150 mM).

3. Methods 3.1. Polyelectrolyte Multilayer (Option 1) 1. Rinse PDMS channels with 0.1 M NaOH for 4 min, then water for 4 min by filling inlet reservoirs with the rinsing solution and applying a vacuum to the waste reservoir (see Note 1). 2. Fill the channel with 5% PB solution to form the cationic layer and let stand 2 min, then remove with vacuum. 3. Let stand 15 min empty (see Note 2). 4. Fill the channel with 3% DS solution to form the anionic layer and let stand 2 min, then remove with vacuum. 5. Let stand 15 min empty. 6. A subsequent cationic layer can be applied if desired by repeating steps 2 and 3 (see Note 3).

3.2. Polyelectrolyte Multilayer (Option 2) 1. 2. 3. 4. 5. 6. 7. 8. 9.

Separate channel section of device from cover film. Rinse channel section with NaOH at 55°C for 15 min (see Note 4). Rinse with water thoroughly and dry with nitrogen. Pipet PAH solution (cationic) onto device, completely covering channels. Allow to stand for 30 min. Remove the PAH solution by rinsing thoroughly with water. Pipet PSS solution (anionic) onto device, completely covering channels. Allow to stand for 30 min. Continue to alternate the PAH and PSS solutions for 5 min with water rinses between each deposition, 14–15 layers should be present depending on the desired nature of the top layer (cationic or anionic) (see Note 5). 10. Cover with the PDMS film and cut inlet and outlet wells using a cork bore.

3.3. UV Graft Polymerization 1. Clean and thoroughly dry PDMS microfluidic device segments (channels and cover film). 2. Prepare an aqueous solution of 0.5 mM NaIO4, benzyl alcohol (0.5% wt), and one of the monomers listed in Subheading 2.3. (10% wt). 3. Immerse the device segments in the monomer solution. 4. Place the immersed segments under a 200-W mercury lamp, with the distance between the sample and the lamp at 5 cm. 5. Irradiate the segments while immersed in the monomer for 3.5 h, rotating the container to ensure even UV irradiation. 6. Remove the segments to distilled water at 80°C and wash under constant stirring for 24 h to remove excess monomer and polymer. 7. Dry under vacuum at room temperature.

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3.4. PDMS Coating on Glass Microchips 1. Prepare a 1% (v/v) PDMS solution in dried and distilled dichloromethane. 2. Load 0.1 M NaOH solution into the inlet well of the microchip. Draw the NaOH solution into the channel using a vacuum. Let solution stand in the channel for 30 min. 3. Rinse repeatedly with purified water by loading water in the inlet well and drawing through the channel with vacuum. 4. Thoroughly dry the channel with dry N2 gas. 5. Rinse the channel with dichloromethane repeatedly using vacuum. 6. Apply the PDMS coating by loading the 1% PDMS solution into the channel and allowing it to stand in the channel for 10 min. 7. Remove the solution using a vacuum. 8. Flush the remaining excess PDMS using dry N2 gas. 9. Place the coated microchip in an oven, increasing the temperature 10°C/min until the temperature reaches 400°C. Allow the chip to remain in the oven at 400°C for 0.5 h. 10. Cool to room temperature in the oven (see Note 6).

3.5. Amine-Terminated PMMA 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Clean the PMMA surface by first soaking in 2-propanol for 15 min. Rinse thoroughly with 18 MΩ⋅cm water. Dry the PMMA pieces under dry N2 gas. Place PMMA pieces in a sealed vessel and purge for 20 min with dry N2 gas. Using a syringe, transfer N-LDAP onto the PMMA pieces without corrupting the N2 environment. Allow the N-LDAP to react with the PMMA for 10 min. Add enough water to completely cover the PMMA pieces to quench the reaction between the PMMA and the N-LDAP. Remove from the reaction flask and rinse the aminated surface thoroughly with purified water. Dry with N2 gas. Place freshly amine-terminated PMMA in an airtight vessel and purge with nitrogen for 20 min. For subsequent attachment of the octadecane: a. Dispense n-octadecane-1-isocynate onto the PMMA without corrupting the N2 environment. b. Allow the PMMA to remain exposed to the n-octadecane-1-isocynate for 10 min. c. Remove the PMMA pieces and quickly rinse with copious amounts of hexanes, toluene, and then acetone. d. Dry with N2 gas. e. Bond the top and bottom PMMA sections thermally (see Note 7).

3.6. Immobilization of Biotin Within a Phosphotidylcholine Bilayer 1. Thoroughly clean and dry PDMS channels and the glass or PDMS cover sheet. 2. Render all PDMS surfaces hydrophilic by exposing to oxygen plasma treatment for 15 s (PDC-32G plasma cleaner, Harrick Scientific, Ossining, NY) (see Note 8).

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3. Prepare vesicles by dissolving the lipids in chloroform and combining them in the mole ration of 0.1% mol biotinylated lipids to DLPC. 4. Evaporate the chloroform in a stream of dry nitrogen, then vacuum evaporate for 4 h. 5. Reconstitute the dried lipids in pH 7.4 phosphate buffer, then sonicate for 5 min using a titanium tip. 6. Centrifuge the solution at 94,500g for 30 min. 7. Centrifuge the supernatant at 176,900g for an additional 3 h. 8. Using a syringe pump (Harvard PHD 2000) infuse the vesicles into the channels and allow to fuse for 30 min. 9. Flush excess vesicles from the channels using phosphate buffer (pH 7.4). 10. Before introduction of streptavidin conjugates, inject BSA solution and incubate for at least 1 h to ensure passivation of any defect sites in the bilayer coating. 11. Wash excess BSA from the channels using pH 9.8 buffer solution.

4. Notes 1. Filling and extraction of PDMS channels is best performed with vacuum, as positive pressure may overwhelm the adsorptive forces bonding the PDMS to a support substrate. For channels that are 50 µm or less, a disposable syringe can be used to create this pressure. Cut a small piece of tubing to fit the end of a syringe (with no needle attached). Heat the end of a metal bolt or nail that is slightly larger than the tubing. Work the end of the tubing onto the end of the bolt creating a flange at the tubing end that is slightly larger than the inlet or outlet well opening. To create the necessary pressure, seal the flanged end of the syringe and tubing over the outlet well and draw the fluid through the channel. The fluid can be removed from the channel using a glass capillary or the end of a filter paper. 2. Though this procedure does not call for a water rinse between PEM coatings, it may be necessary to perform a rinse at this juncture to improve uniformity of the coatings. 3. Atomic force microscopy is useful to verify the PEM coating. For an immediately tangible test, the wettability of the channel is also useful in determining if the coating has adsorbed properly. 4. Do not rinse the PDMS cover film with NaOH. 5. Before the cover film is applied, you can check the efficacy of the coating using contact angle measurements. 6. Addition of 0.10% Tween-20 to your separation buffer will provide well-defined peaks for protein samples. 7. Thermal bonding in this procedure was accomplished by heating each surface of the PMMA on a hotplate to 150°C for 5–10 min. Do not heat the entire PMMA part in a furnace, as outgassing and bubbling are possible. Align the two device pieces, preferably in an alignment holder, then place a 50 lb weight on the PMMA pieces and allow to cool to room temperature slowly in a programmable oven over 2 h. 8. Oxygen plasma treatment of the surfaces for 1 min should be sufficient. Once treated, the PDMS will bond more strongly adsorptively and can form covalent bonds with the glass substrate. Because of this, the PDMS channels cannot easily be removed after this step.

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References 1. Badal, M. Y., Wong, M., Chiem, N., Salimi-Moosavi, H., and Harrison, D. J. (2002) Protein separation and surfactant control of electroosmotic flow in poly(dimethylsiloxane)-coated capillaries and microchips. J. Chromatogr. A 947, 277–286. 2. Barker, S. L. R., Tarlov, M. J., Canavan, H., Hickman, J. J., and Locascio, L. E. (2000) Plastic microfluidic devices modified with polyelectrolyte multilayers. Anal. Chem. 72, 4899–4903. 3. Gottschlich, N., Jacobson, S. C., Culbertson, C. T., and Ramsey, J. M. (2001) Twodimensional electrochromatography/capillary electrophoresis on a microchip. Anal. Chem. 73, 2669–2674. 4. Henry, A. C., Tutt, T. J., Galloway, M., et al. (2000) Surface modification of poly(methyl methacrylate) used in the fabrication of microanalytical devices. Anal. Chem. 72, 5331–5337. 5. Hu, S. W., Ren, X. Q., Bachman, M., Sims, C. E., Li, G. P., and Allbritton, N. (2002) Surface modification of poly(dimethylsiloxane) microfluidic devices by ultraviolet polymer grafting. Anal. Chem. 74, 4117–4123. 6. Katayama, H., Ishihama, Y., and Asakawa, N. (1998) Stable cationic capillary coating with successive multiple ionic polymer layers for capillary electrophoresis. Anal. Chem. 70, 5272–5277. 7. Liu, Y., Fanguy, J. C., Bledsoe, J. M., and Henry, C. S. (2000) Dynamic coating using polyelectrolyte multilayers for chemical control of electroosmotic flow in capillary electrophoresis microchips. Anal. Chem. 72, 5939–5944. 8. Eteshola, E. and Leckband, D. (2001) Development and characterization of an ELISA assay in PDMS microfluidic channels. Sens. Actuators B Chem. 72, 129–133. 9. Mao, H. B., Yang, T. L., and Cremer, P. S. (2002) Design and characterization of immobilized enzymes in microfluidic systems. Anal. Chem. 74, 379–385.

6 Polyelectrolyte Coatings for Microchip Capillary Electrophoresis Yan Liu and Charles S. Henry Summary In chip-based electrophoretic analysis of biomolecules, chemical modification of the microchannel is widely employed to reduce or eliminate the analyte–wall interactions and alter electroosmotic flow (EOF) in the microchannel. A stable polyelectrolyte multilayer coating is one common way to regulate or eliminate EOF and prevent analyte adsorption for the rapid, efficient separation of biomolecules within microchannels. A wide variety of polyelectrolytes have been used as coatings. This chapter deals with how to coat microchips with polyelectrolytes and the expected results using polybrene and dextran sulfate as models. The technique presented here is generally applicable to any polyelectrolyte. Key Words: Polyelectrolyte; coating; microchip; capillary electrophoresis; electroosmotic flow; poly(dimethylsiloxane); polybrene; dextran sulfate.

1. Introduction Recently, microchip capillary electrophoresis (CE) has become a powerful tool to analyze biomolecules (1,2). Initially, work focused on microfabrication of glass microchips because of the mature micromachining technology (3,4). However, the time- and labor-consuming fabrication process of glass microchips and the requirement of clean-room facilities have led to the investigation of alternative substrate materials for the construction of CE microchips (5,6). Poly(dimethylsiloxane) (PDMS) attracted significant attentions as a microchip substrate because of its potential for mass production, rapid protyping, and good optical properties. However, peak tailing owing to the analyte absorption into PDMS microchip has been well documented for nonpolar hydrophobic species (7,8). In addition, one significant problem for PDMS devices is poorly defined electroosmotic flow (EOF) (9). Under CE conditions, the EOF dominates the flow velocity of both the run buffer and the analytes being separated. From: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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In PDMS microchips, the nature of EOF is dependent on the process used for sealing chips, which generates different surface charge densities (9,10). In addition, the use of multiple substrate materials in a single device may cause inconsistency of the flow velocity and diminish separation efficiency owing to nonuniform flow in the capillary columns resulting from the different ζ-potentials of different substrate materials. Generally, these discrepancies in PDMS substrates are the result of minimal characterization of surface ionizable groups under typical CE conditions. Moreover, the EOF decreases with pH, making rapid separations of mixtures of anions and cations difficult at low pH. To overcome these problems, several surface modification techniques, including dynamic, covalent, and noncovalent coating, have been used to control the EOF (11–13). Dynamic coating is typically prepared by rinsing the capillary with a solution containing a coating agent that is either a polymer or a small molecular-mass inorganic ion. A small amount of coating agent is also added to the run buffer to keep the coating on the capillary wall surface. The life time of dynamic-coated capillaries can be extended by using an occasional, simple regeneration process. The resulting coating is used for both EOF suppression and prevention of protein adsorption. Covalent coating is the most prevalent, and perhaps the most effective strategy to control EOF and prevent biomolecule adsorption. Covalent coating involves the covalent immobilization of molecules onto the capillary wall. However, lengthy derivatization procedures, unstable coating layers beyond a limited pH range, and poor reproducibility limit the wide use of covalent coatings. To overcome the drawbacks associated with covalent derivatization, charged polyelectrolytes, such as polybrene and polyethyleneimine, have been used for adsorbed noncovalent capillary coatings. Owing to the strong electrostatic attraction between these polycations and the anionic silanols on the capillary inner surface, these polymers adsorb strongly onto the capillary inner wall. The capillary can be coated, regenerated, and then recoated, making them more cost and time effective than covalent coatings. 2. Materials 2.1. Equipment 1. 2. 3. 4.

Harrick plasma cleaner/sterilizer PDC-32G. Fisher FS20 sonicator. Voltameter. High voltage power supply (Standford PS350/5000V-25W, or Spellman CZE 1000R or similar). 5. Whatman 0.2 µm-syringe filter. 6. 1 kΩ resistor. 7. Two platinum wire electrodes (diameter: 0.5 mm).

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2.2. Chemicals and Reagents 1. 2. 3. 4. 5. 6.

Sylgard 184 silicone elastomer and curing agent. Polybrene. Dextran sulfate. Phosphate buffers: 20 mM pH 3.0–10.0. Sodium hydroxide. Methanol.

3. Methods 3.1. PDMS Device Fabrication A brief summary of the fabrication of PDMS microchips is provided here. A more detailed discussion of PDMS fabrication is presented in the appropriate chapter in this text (Chapter 3). A degassed mixture of sylgard 184 silicone elastomer and curing agent (10:1) was poured onto a 3-in. silicon mold that was patterned by lithographic technique and had been cleaned sequentially with deionized water (DI) and methanol and dried with a stream of nitrogen gas. After at least 2 h of curing at 65°C, the PDMS replica was peeled from the mold, resulting in a pattern of negative relief channels and reservoirs in the PDMS. Buffer reservoirs were then opened with a circular punch and the PDMS was trimmed to size with a razor blade. Bare PDMS replicas were formed by casting the PDMS mixture on a dry, clean nonpatterned silicon wafer. 3.2. Microchip Sealing Modifications of previously published reversible and irreversible sealing methods can be used to assemble the microchips (9,10). Reversible sealing involved thoroughly rinsing a PDMS replica and a glass plate (or a second piece of PDMS) with methanol and bringing the two surfaces into contact with one another prior to drying. The assembled microchip is then dried in an oven at 65°C for 10 min. The air bubbles between the two layers are driven out by pressure. This method of reversible sealing gives the most consistent sealing and does not require the use of clean room/hood facilities. Irreversible sealing is accomplished by first thoroughly rinsing a PDMS replica and a glass plate with methanol, and then drying them separately under a stream of nitrogen. The two pieces are then placed in an air plasma cleaner and oxidized at high power for 45 s. The substrates are brought into conformal contact immediately after removal from the plasma cleaner and an irreversible seal forms spontaneously. This seal is sufficiently strong that the two surfaces can not be separated without destroying the assembled microchip. 3.3. Noncovalent Coating Although channels can be coated with different polyelectrolyte layers, the whole coating procedure is similar to the one that was developed by Katayama

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Fig. 1. Schematic for multilayer coating. (A) Preconditioned native poly(dimethylsiloxane) microchip, (B) first layer coating with a 5% polybrene water solution, and (C) second layer coating with 3% dextran sulfate water solution.

and coworkers for the conventional CE (14,15) and is applicable to microchips constructed from a variety of materials. Briefly, the channel is preconditioned for a few minutes then sequentially flushed with the desired polyelectrolyte solutions. A schematic of the whole procedure is shown in Fig. 1, with polybrene (PB) and dextran sulfate (DS) coating the channel as an example. Briefly, the separation channel is rinsed with 0.1 M NaOH for 4 min and flushed with DI for 4 min, respectively. Once preconditioned, the channel is sequentially filled with 5% PB solution and 3% DS solution (both in water) for 2 min each with a 15-min waiting period after each rinse. This procedure of successive coating results in a bilayer of PB/DS on the channel walls. The polymer rinsing steps can be repeated multiple times to build up additional layers. Finally the channel is flushed with the run buffer solution.

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3.4. EOF Measurements It is important to measure the EOF to ensure that an appropriate coating has been generated. For more detailed information on measuring EOF consult the appropriate chapter (see Chapter 13). Briefly, the running electrolyte for electrophoresis experiments is 20 mM pH 3.0–10.0 phosphate buffer. The pH was established by titrating a solution of either o-phosphoric acid or sodium dihydrogen phosphate with sodium hydroxide. All buffers are prepared in DI, passed through a 0.20-µm pore size syringe filter, and degassed for 5 min in a sonicator before use. A modification of a previously published current monitoring method is used to determine the EOF (16). Both reservoirs are filled with dilute buffer (2:1 buffer: water), and the channels are subsequently conditioned under an electric field of 1200 V for 15 min. The increased dilution factor as compared with standard protocol (19:1 buffer: water) is used to ease end point detection. No statistical differences in the absolute values are noted between the two protocols. A 1 kΩ resistor is placed in line between the waste reservoir and electrical ground to follow the separation current. A voltameter records the potential changes across the resistor, which correlates to the current through Ohm’s Law. The sample reservoir is then filled with concentrated buffer, and the potential is reapplied. The time required for the current plateau is measured for each run and is indicative of the concentrated buffer’s filling the separation channel. The sample reservoir is then filled with dilute buffer and the above procedure repeats. The time required for the current to reach this plateau was used as the migration rate of a neutral marker, and the EOF is determined by µEOF = L2 / Vt

(1)

where L is the length of the separation channel, V is the total applied voltage, and t is the time in s required to reach the new current plateau. The typical voltage profile across the resistor is shown in Fig. 2, and the time to reach a current plateau is 53 s for this microchip. This is a modification of the traditional mobility equation that takes into account that the total and effective capillary lengths are identical. 3.5. Stability and Reproducibility of the Coating Layer One concern with the noncovalent coating is the stability of the coating. A pH of 3.0 is chosen for evaluation of the PB-coating lifetime because no EOF is detected at this pH point for native PDMS microchips. As the coating becomes detached from the channel wall, the EOF will approach zero until the coating becomes completely detached, at which point the EOF direction will be reversed and no current change will be detected. Another concern with the noncovalent coating is the reproducibility of the coated layer from chip-to-chip. The EOF is detected in six PB/DS-coated

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Fig. 2. Typical voltage profile for electroosmotic flow measurement. Table 1 Comparison of Electroosmotic Flow for Uncoated, PB/DS-Coated, and PB-Coated PDMS/Glass Microchipa pH Uncoated chip PB/DS-coated chip PB-coated chip aUnit:

3.0 0 2.47 –4.29

4.0 2.73 2.93 –3.34

5.0 3.06 3.29 –2.40

6.0 3.01 3.63 –2.95

7.0 4.04 3.65 –2.21

8.0 3.89 3.63 –1.93

9.0 4.43 3.63 –1.97

10.0 4.89 3.69 –1.95

× 10–4 cm2⋅V–1⋅s–1.

PDMS/glass chips at three different pH values in different days. The relative standard deviation (RSD) of the EOF for six PB/DS-coated PDMS/glass chips is considered as an indicative parameter of the complete and effective covering of the channel walls by the PB and DS layers. 4. Notes 1. Although the EOF differs significantly between uncoated oxidized and native PDMS/glass microchips, the PB/DS coating layer can compensate for the differences in type and density of anionic groups on the surface, as well as the difference between chip-sealing techniques, and generates a constant EOF regardless of substrate and sealing technique. The coating is solely responsible for the generation and control of EOF in the microchannels. The EOF results from native-, PB-, and PB/DS-coated microchips are shown in Table 1. 2. If electrochemical detection is involved in the microchip CE, one concern is the electrode fouling resulting from either analyte or polymer adsorption on the work-

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ing electrode surface. This is of particular interest when using a coated microchip in which any detached coating could potentially adsorb onto the electrode surface, resulting in the decrease of detection signal. Separations of dopamine and hydroquinone in both PB/DS-coated and uncoated microchips can give the demonstration whether the detached coating adsorb on the working electrode surface. 3. The noncovalent-coating technique is applicable to numerous polyelectrolytes, among which polybrene is the most popular and is sold commercially as a capillary treatment. In addition, polyamine, poly(dimethyl diallyl ammonium chloride), polyethyleneimine, and polyarginine are widely used to prevent biomolecule adsorption on the capillary surface.

References 1. Barta, C., Ronai, Z., Nemoda, Z., et al. (2001) Analysis of dopamine D4 receptor gene polymorphism using microchip electrophoresis. J. Chromatogr. A 924, 285–290. 2. Fanguy, J. C. and Henry, C. S. (2002) Pulsed amperometric detection of carbohydrates on an electrophoretic microchip. Analyst 127, 1021–1023. 3. Culbertson, C. T., Jacobson, S. C., and Ramsey, J. M. (2000) Microchip devices for high-efficiency separations. Anal. Chem. 72, 5814–5819. 4. Kopp, M. U., Mello, A. J., and Manz, A. (1998) Chemical amplification: continuousflow PCR on a chip. Science 280, 1046–1048. 5. Martin, R. S., Gawron, A. J., and Lunte, S. M. (2000) Dual-electrode electrochemical detection for poly(dimethylsiloxane)-fabricated capillary electrophoresis microchips. Anal. Chem. 72, 3196–3202. 6. McClain, M. A., Culbertson, C. T., Jacobson, S. C., and Ramsey, J. M. (2001) Flow cytometry of Escherichia coli on microfluidic devices. Anal. Chem. 73, 5334–5338. 7. McDonald, J. C., Duffy, D. C., Anderson, J. R., et al. (2000) Fabrication of a configurable, single-use microfluidic device. Electrophoresis 21, 27–40. 8. Effenhauser, C. S., Bruin, G. J., and Paulus, A. (1987) Integrated chip-based capillary electrophoresis. Electrophoresis 18, 2203–2213. 9. Ocvirk, G., Munroe, M., Tang, T., Oleschuk, R., Westra, K., and Harrison, D. J. (2000) Electrokinetic control of fluid flow in native poly(dimethylsiloxane) capillary electrophoresis devices. Electrophoresis 21, 107–115. 10. Duffy, D. C., McDonald, J. C., Schueller, O. J. A., and Whitesides, G. M. (1998) Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70, 4874–4884. 11. Giordano, B. C., Copeland, E. R., and Landers, J. P. (2001) Towards dynamic coating of glass microchip chambers for amplifying DNA via the polymerase chain reaction. Electrophoresis 22, 334–340. 12. Badal, M. Y., Wong, M., Chiem, N., Salimi-Moosavi, H., and Harrison, D. J. (2002) Protein separation and surfactant control of electroosmotic flow in poly(dimethylsiloxane)-coated capillaries and microchips. J. Chromatogr. A 947, 277–286. 13. Horvath, J. and Dolnik, V. (2001) Polymer wall coatings for capillary electrophoresis. Electrophoresis 22, 644–655.

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14. Katayama, H., Ishihama, Y., and Asakawa, N. (1998) Stable cationic capillary coating with successive multiple ionic polymer layers for capillary electrophoresis. Anal. Chem. 70, 5272–5277. 15. Katayama, H., Ishihama, Y., and Asakawa, N. (1998) Stable capillary coating with successive multiple ionic polymer layers. Anal. Chem. 70, 2254–2260. 16. Huang, X., Gordon, M. J., and Zare, R. N. (1988) Current-monitoring method for measuring the electroosmotic flow rate in capillary zone electrophoresis. Anal. Chem. 60, 1837, 1838.

III DETECTION METHODS FOR MICROCHIP CAPILLARY ELECTROPHORESIS

7 Interfacing Microchip Capillary Electrophoresis With Electrospray Ionization Mass Spectrometry Trust Razunguzwa and Aaron T. Timperman Summary Microfluidic devices are a unique enabling technology for chemical separations, modification, and synthesis that are ideally suited for the manipulation of low volume samples on the order of a few nanoliters in volume. Complex patterns of capillary-sized channels with zero dead volume connections are the distinguishing features of many microfluidic devices. Concurrently, mass spectrometry has undergone further development, and is now arguably the method of choice for structural characterization of mass- and volume-limited samples. The production of ions in the gas phase from the solution phase is critical for direct coupling of fluidic devices with the mass spectrometer, and the electrospray ionization (ESI) sources are well suited for this application. Micro- and nanoflow ESI interfaces are ideal for these applications as they cover flow rate ranges from the hundreds to a few nanoliters per minute, which are the same as the flow rates used by most microfluidic devices. Herein, the assembly and operation of a simple ESI interface for coupling a microfluidic device and mass spectrometer is described. Key Words: Capillary electrophoresis; microfluidic device; mass spectrometry; electrospray; interface.

1. Introduction Microfluidic devices have a promising future for chemical analysis and synthesis at low volume and mass, while being amenable to massive parallelism for increasing sample throughput. Functionalities, which have been developed on the chip, include sample preparation, separation of complex mixtures, preconcentration of analytes, and tryptic protein digestions (1–4). The ability of microfluidic devices to handle and manipulate extremely small volumes of solutions, in the nanoliter regime, make them especially attractive for biological samples whose amounts are frequently limited. Fluidic networks are characterized by intersections with zero dead volume enabling efficient processing of complex samples. In general, microchips for chemical analysis allow parallel From: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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sample analysis, shorter analysis times, increased separation efficiencies, lower detection limits, and reduced reagent consumption and waste generation (2,5). The mass spectrometer is a powerful analytical tool owing to its high resolving power, ion isolation capability, and ability to measure the mass to charge ratio of both parent and fragment ions. Electrospray ionization mass spectrometry (ESI-MS) is concentration (not mass) sensitive. Therefore, miniaturizing the sample introduction system achieves the highest sample concentrations and the lowest detection limits. Microfluidic devices interfaced with ESI-MS provide a convenient, miniaturized sample preparation/introduction system. Solutions are commonly transported through microfluidic channels by application of an electric potential (electrokinetically) or by applying a pressure (hydrodynamically). In addition, flow control on microfluidic devices can also be governed electrokinetically, although the development of reliable mechanical valves is desirable. Most systems interfaced with mass spectrometers are electrically driven systems, and these will be our primary focus (6–8). Early attempts to interface microchips to MS involved spraying fluid directly from an exposed channel on the microchip (9,10). Although this design was attractive, in that the design did not require complex machining, it resulted in large dead volumes owing to the formation of large Taylor cones from the solution exiting out of the open end of the microchannel, leading to sample dilution (lower sensitivities) and band broadening. Also, because the surface at the edge of the microchip was flat, these devices required an impractically high voltage to overcome the liquid surface tension and initiate electrospray. Other researchers attempted using hydrophobic coatings (to minimize surface tension) at the edge of the microchip and on-chip nebulizers, but this has been met with limited success (11). More recently, spraying from a transfer capillary spray tip attached to a microchip has been found to be an effective way of sample delivery into a mass spectrometer, as it provides a spray tip from which electrospray can be more easily generated (12–14). For these devices, application of high voltage can be through a platinum electrode inserted in solution, liquid junction, or by a conductive coating at the outlet of the capillary. Most recent advancements are utilizing microfabrication techniques to integrate nanospray tips directly onto the microfluidic device, which eliminates band broadening associated with the dead volume at the chip–capillary interface and the extra column volume associated with capillary spray tip-coupled microfluidic devices. An example is a device developed by the Henion group and Advion Biosciences that is based on a silicon substrate with etched nozzles in the planar surface of the silicon wafer, used directly as microspray emitters perpendicular to the chip (15). Smith’s group also recently reported a polycarbonate-based device with an ESI tip constructed on polycarbonate plates by laser micromachining for isoelectric focusing-ESI-MS (16).

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Considering the different ways in which a microchip–MS interface can be configured, it is of paramount importance to identify the parameters that affect electrospray generation at the outlet of the microchannel or capillary to obtain a functional interface. The first requirement for a chip–MS interface is maintaining bulk flow of solution to the spray tip, which is needed to sustain droplet formation in the electrospray source (10). For an electrokinetically driven system, this means that electroosmotic flow (EOF) must be present in the microchannel, requiring a high density of charged sites on the channel surface. A basic pH is usually sufficient to meet this requirement given that many microfluidic devices are fabricated on glass with silanol groups that deprotonate as pH increases. Stable spray is dependent on the diameter of the tip and the spray capillary inner diameter (I.D), which generate more resistance to flow as they get smaller. At the low flow rates afforded by microfluidic devices, small spray tip sizes and spray capillary I.Ds, small droplets with high surface-to-volume ratio form at the tip, giving rise to better ionization and desolvation efficiencies because of rapid droplet solvent evaporation. Mann and Wilm estimated the droplet size formed at a 1- to 2-µm tip to be less than 200 nm in diameter, corresponding to an average concentration of one analyte molecule per droplet for a 1-pmol/µL solution (17). By separating molecules into different droplets, cluster formation is minimized and improves analysis of sample with high-salt concentrations. McLafferty et al. demonstrated this by further reducing the flow rate of 25 nL/min used by Mann and Wilm to 1 nL/min using a 5-µm I.D, 2-µm spray tip, and achieved attomole sensitivity detection of large biomolecules (18). It is, therefore, beneficial to use spray capillaries with the smallest capillary I.Ds and spray tip sizes to achieve efficient electrospray and consequently high sensitivities. Other factors need to be considered for the operation of a microchip–MS interface. The lowest detection limits in ESI-MS are usually realized when the ionization source is operated in the positive ion mode (i.e., a positive voltage is applied at the spray tip). This requirement dictates a low pH spray solution. However, in order to support sufficient EOF under acidic conditions, the native glass surface of the microchannel must be coated with a material that provides a high surface charge at acidic pH. A make-up or sheath flow solution may be necessary in some cases to modify the electrophoresis buffer to achieve the necessary conditions (pH, concentration, or flow) for electrospray. Finally, the chip substrate (usually glass) must be compatible with organic solvents and acids while producing minimal chemical background. Solvents must be of highest obtainable purity and volatile without ion suppressing agents such as triflouroacetic acid (TFA). The assembly of the chip capillary electrophoresis (CE)–ESI-MS interface starts with fabrication of the microfluidic device using standard photolithography and wet chemical etching procedures to introduce predesigned

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microchannels onto the glass surfaces (19). Compared to the use of polymers, glass is preferable because of its excellent optical properties and its stability toward organic solvents. Bonding to enclose the channels can be achieved with direct bonding at low and high temperature without adhesives. This is particularly important for chip ESI-MS detection because fillers, unreacted monomer, and plastic can increase the background signal. This process is followed by surface coating of the channels with a positively charged coating to increase EOF and minimize analyte adsorption. Reservoirs are attached to the chip, and the chip is conditioned with buffer followed by spray tip attachment. Finally, the microchip is brought in front of the MS for sample introduction. 2. Materials 2.1. Equipment 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

Sawed/scribed wafer cleaner (Ultra and Equipment Co., Fremont, CA). Airclean 600 workstation (Airclean Systems, Raleigh, NC). Blue tape and mounting rings for holding glass plates in wafer washer. DC power supply (Agilent, Palo Alto, CA). High voltage power supply (Matsusada Precision Inc., San Francisco, CA). Stereomicroscope GZ6 (Leica Microsystems, Bannockburn, IL). Multimeter (Fluke, Everett, WA). Electrospray ionization mass spectrometer (Thermo Finnigan, San Jose, CA). Soda lime glass plates with chromium layer and photoresist (Telic Co., Santa Monica, CA). Ultraviolet (UV) lamp and aligner (Advanced Radiation Corporation, Santa Clara, CA). Stylus instrument (Tencor Instruments, Milpitas, CA). Fused silica capillary (Polymicro Technologies, Phoenix, AZ). High precision drill press (Tralmike’s-Tool-A-Rama, Plainfield, NJ). Carbide microdrills (Kyocera Tycom, Owego, NY). Nanoport assemblies (Upchurch Scientific, Oak Harbor, WA). Three-dimensional translational stage (Newport). Laser puller (Sutter Instruments, Phoenix, AZ). Syringe pump (Harvard Apparatus, Holliston, MA).

2.2. Reagents 1. Glass etching solution: 50% hydrofluoric acid (HF)/60% nitric acid/water, (2/1/7 v/v/v). 2. Piranha solution: concentrated sulphuric acid/30% hydrogen peroxide, (3/1 v/v). 3. Ammonium hydroxide reagent: concentrated ammonium hydroxide/30% hydrogen peroxide/water, (1/1/5 v/v/v). 4. Acetone. 5. 10% Tetramethylammonium hydroxide developer solution (Clariant).

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Fig. 1. An outline of the photolithography, wet chemical etching, and bonding processes of glass microchips. 6. Soap solution: one scoop of powdered soap in 1 L of water. 7. Make-up solution and separation buffer: 0.1% acetic acid and 5% acetonitrile in water. 8. N-trimethoxypropyl-N,N,N-trimethyl ammonium chloride (TMAC) (Gellest). TMAC is toxic and has a very strong odor.

3. Methods 3.1. Chip Fabrication Fabrication of glass microchips is carried out using standard photolithography and wet chemical etching (20) on a 10 × 5 cm soda-lime glass (see Note 1), followed by drilling of access holes on the glass substrate, and finally, lowtemperature and high-temperature bonding of the etched glass plate to a cover plate (21). An outline of the standard fabrication process from photolithography to bonding is shown in Fig. 1. Glass plates coated with chromium and photoresist layers are commercially available. 3.1.1. Photolithography 1. Channel patterns are designed and drawn using Macromedia Freehand 10. Negative transparencies of these patterns are printed on an Afga Accuset 1000 printer with a resolution of 3000 dpi and used as photomasks in the photolithography process.

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2. A soda-lime glass chip with photoresist is placed on the aligner and held in place by vacuum. The photomask with the channel pattern is aligned and placed over the glass chip. 3. The chip and mask are positioned under the UV lamp and exposed for 15–40 s at 365 nm depending on the smallest channel size on the mask (see Note 2). 4. After exposure to UV light, the photoresist is developed. The photoresist polymer that was exposed to UV light is washed away by the developer solution AZ 312 MIF/H2O (1:1), leaving the pattern of the photomask on the photoresist layer. 5. The channel pattern on the photoresist layer is inspected using a microscope to check for poorly developed channels.

3.1.2. Wet Chemical Etching 1. A chromium etchant solution is used to remove the chromium within the channel pattern leaving an exposed glass layer. The chips are placed in the chromium etchant solution for 2 min. The glass chips are then rinsed with deionized (DI) water and blown dry with nitrogen. 2. The photoresist layer is stripped using acetone and the channels are inspected for completeness of the chromium etching using the microscope. 3. The glass substrates are etched with 50% HF/60% HNO3/H20, (2/1/7 v/v/v), rinsed with DI water, and blown dry with acetone. Caution: HF should be handled with extreme care as it is very hazardous. Exposure to vapor and skin contact is extremely dangerous and, therefore, the accepted methods for handling and use should be followed. The chromium layer is removed using the chromium etchant solution after glass etching and DI water rinsing of the glass plates (heat to 50°C if necessary) (see Note 3). 4. The chip is rinsed with DI water and blown dry with nitrogen gas soon after the channels are inspected for completeness using a microscope. Channel profiles and dimensions are then obtained using the stylus instrument (see Note 4).

3.1.3. Glass Bonding The glass bonding procedure is outlined next: 1. Clean the glass plates (etched plate and cover plate) with acetone using a lint free swab. 2. Clean the glass plates with detergent solution (one scoop of powdered soap in 1 L of DI water) and rinse with DI water. 3. Place the glass plates in piranha solution at 100°C for 45 min to 1 h and rinse with DI water. 4. Place the glass plates in ammonium hydroxide reagent at 80°C for 45 min to 1 h and rinse with DI water. 5. Wipe down the Airclean workstation with a lint-free cloth containing a few squirts of isopropanol. Clean again with mitten if necessary. Be sure to wipe down all the parts to the washer. Wipe down mounting rings and place blue tape onto two mounting rings.

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Fig. 2. (A) Illustration of the vertical access hole drilled at the end of a channel on the microchip. (B) Shows a minimal dead volume connection of a fused silica capillary spray tip to the microchip. The 360- and 650-µm holes are drilled using carbide microdrills. A silica seal-tight sleeve is inserted into the 650-µm hole and holds a 360-µm O.D. capillary spray-tip, which extends into the 360-µm hole. There is a tight seal between the sleeve and the 650-µm hole, and between the 360-µm spray-tip and the 360-µm hole (see Note 8). 6. Load the glass plates on the blue tape and place them in the high-pressure washer and perform one washing and one drying cycle. 7. Align the glass plates using a spacer ring in between the mounting rings and bond them using a rubber roller to press them together. Eliminate all Newton rings by squeezing them out from in between the glass plates. Be sure to bond the etched side to the plain glass plate (see Note 5). 8. Permanent bonding is achieved by placing the glass plates in the furnace and applying the temperature program: 25–100°C at 40°C/min, 100°C for 15 min, 100–600°C at 10°C/min, 600°C for 15 min. The furnace was allowed to cool naturally to ambient temperature.

3.1.4. Access Hole Drilling The access holes to the channels of the microchips are drilled directly on the etched glass plate before bonding using a high-precision drilling press and carbide microdrills ranging from 200 to 650 µm. As shown in Fig. 2A, the etched glass plate is first drilled half way (0.5 mm) through the glass from the open channel side using a 200-µm microdrill, and then from the other side using a 360-µm microdrill until it meets the 200-µm hole drilled from the other side of the glass (see Note 6). The bottom of the 360-µm hole is flattened using a flat endmill microdrill. This drilling method of the access holes allows for the tight fit connection of a 360-µm outer diameter (O.D) fused silica capillary to the

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microchip with its end lying flat at the bottom of the 360-µm hole and the capillary bore emptying into the 200-µm hole. The drilling process is carried out with the aid of a microscope to align the microdrill to the end of a channel as well, as aligning the microdrill to the hole drilled from one side halfway through the glass, when drilling from the other side. A microscope is also used to follow the progress of the drilling process. The connection of the spray tip to the microchip is achieved by drilling microholes through the junction between the cover plate and the etched glass plate, at the edge of the bonded microchip. A 360-µm hole is first drilled 8 mm into the glass chip through a channel running off the edge of the microchip. A 650-µm hole is then drilled through the 360-µm hole 5 mm into the glass plate before an end mill microdrill is used to flatten the bottom (see Note 7). Figure 2B shows the schematic cross-section of the end result of this drilling process. 3.1.5. Microchip Fittings Polyetheretherketone (PEEK™) polymer Upchurch nanoport assemblies are attached to the microchips via preformed adhesive rings to act as reservoirs leading to the access holes and connecting fused silica capillaries to the microchip. Two types of nanoport assemblies are used to attach a fused silica capillary column directly to the microchip and these are the 6.32 flat-bottom and 6.32-coned assemblies (Upchurch, F.123S and F.124S, respectively). The Upchurch 1/4.28 FB nanoports are used as reservoirs. 3.2. Microchip Coating A positively charged quaternary amine reagent that provides a positive permanent coating is used to modify the inner surface of the microchip and the fused silica capillary spray tip to give a positively charged surface (see Note 9). The coating generates a strong anodic EOF at low pH, in addition to minimizing adsorption of the positively charged sample components to the channel walls by electrostatic repulsions between the positively charged channel walls and positively charged ions of the sample. The coating procedure is outlined next: 1. 2. 3. 4. 5. 6.

Rinse all the channels with 1 M NaOH for 5 min. Flush DI water through the channels for 3 min. Flush all channels with 10% TMAC and fill all the channels with 10% TMAC. Fill the reservoirs with pure methanol. Allow the chip to stand for 2 h while replenishing evaporated methanol. Wash out unreacted TMAC and the solvent methanol from the channels using DI water followed by the separation buffer.

The coating procedure should be carried out in a fume hood because TMAC is toxic and has a strong offensive odor.

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3.3. Making Spray Tips A fused silica capillary emitter (6 cm, 50 µm I.D., and 360 µm O.D) is attached to the microchip as previously described in Subheading 3.1.4. The spray tips can be made in house with a capillary puller or purchased directly from New Objective (Woburn, MA). Using a laser puller, the emitter exit end is tapered to approx 5 µm I.D. A 15-cm fused silica capillary column is cut and the polyimide coating burnt off the center of the capillary. The section without the polyimide coating is wiped clean using methanol and a Kimwipe. The capillary is placed in the laser puller instrument with the exposed silica portion in the path of the laser. Once the capillary is aligned, the laser is activated and the tip pulled. Two spray emitters are obtained and are cut to the desired length (see Note 10). The laser puller instrument is programmed according to the tip size required and the capillary size. It has been demonstrated, in literature, that emitters with small tip sizes afford better sensitivities than larger ones because they form smaller droplets with high charge densities, which result in better desolvation and ionization efficiencies (15). The spray tips can be etched with 50% HF solution for 30–60 s to further reduce the tip size and thickness of tip wall. 3.4. Interfacing of Microchip to ESI-MS Once all the reservoirs and fittings have been attached to the microchip, the channels are filled with the separation buffer solution by applying a vacuum sequentially to all the reservoirs. It is imperative that no bubbles are introduced into the channels because these can introduce void volumes and current breakdown problems. All the buffers have to be degassed beforehand by sonication or vacuum degassing. The channel resistances are then determined indirectly by measuring the voltage drop across a 10-kΩ resistor to obtain an ohm plot. A high voltage power supply is used to apply voltages to the buffer solutions in the reservoirs. Caution: exercise caution when applying dangerous high voltages to avoid electrocution owing to the large charging capacitance of the power supplies. Good electrical connectivity is tested for all the channels on the microchip by establishing whether a current flows through the electrolyte in the channels when the high voltage is applied. The microchip is mounted on a three-dimensional translational stage and brought in front of the mass spectrometer where the spray tip is aligned to inlet of the MS orifice using the x-y-z manipulator. A distance of approx 3–6 mm is left between the spray tip and the MS inlet orifice. 3.4.1. Microchip ESI-MS The design of the microchip used to interface to ESI-MS is shown in Fig. 3. Reservoirs A, B, and C correspond to the separation buffer, sample, and

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Fig. 3. Schematic representation of the microchip-electrospray ionization mass spectrometry assembly. The microchannels have a positive charge and the device is operated at low pH. Ports A–D are the buffer, sample, waste, and make-up solution reservoirs, respectively. For an electrospray compatible separation buffer, side channel D can be excluded from the design, because in this case no make-up solution is required. Loading of a sample plug is performed by applying a potential difference between buffer reservoirs B and C, with the potential at B being more negative relative to the potential at C. The sample plug is then injected and separated by applying a negative voltage to the separation buffer reservoir A with reservoirs B and C floating. The sample migrates down the separation channel until it reaches the spray tip for the onset of electrospray.

waste reservoirs, respectively. Reservoir D is used to connect to a syringe pump for delivery of the make-up flow solution in case where it is necessary. A sample is dried down and redissolved in the separation buffer followed by the replacement of the buffer solution in reservoir B with the sample solution. The operation of the microchip–ESI/MS interface is outlined as follows: 1. The sample is loaded between the tees by applying a negative voltage to the sample reservoir, ground to the waste reservoir, and leaving reservoir (A) floating (see Note 11). The sample is then injected into the separation channel by applying a negative voltage to the separation buffer reservoir (A) with reservoirs B and C floating and the mass spectrometer at ground (see Note 12). The sample plug migrates down the separation channel with the concomitant separation of the different sample species based on their charge, size, and shape. The channels are coated with a permanent positive coating that generates a strong EOF toward the anode (toward the mass spectrometer) at low pH. The EOF in the microchannel results from the motion of mobile counter-ions in the diffuse region of the electric double layer, in response to the applied electric field. Because the mobile counter-ions in this case are negative ions (for a positively charged wall), the negative ions reach the spray tip first, followed by the neutrals,

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and then the positive ions. The separation of the positive ions is therefore better because they have more time to separate, and these are the ions detected by positive ion mode MS. EOF is usually preferred over pressure-driven flow for onchip separations because of the plug-like flow profile of EOF, which results in better separation efficiencies. However, EOF is not very reproducible because of several factors that can induce velocity gradients in the bulk liquid and these should be minimized. These include partial blockage or narrowing anywhere in the channel, a nonuniform wall surface charge distribution, slight height differences between free surfaces in the reservoirs, capillary forces resulting in curvatures of the free surfaces in the reservoirs, and contact angle hysteresis of a bubble entrapped in the channel. 2. For an electrophoresis buffer compatible with electrospray, the sample bands are delivered to the spray tip by EOF where the electrospray process occurs. Small and charged buffer droplets form at the spray tip and the electric field at the tip generates an electrostatic force that is sufficient to pull the droplets out of the tip toward the ground plate of the mass spectrometer. This phenomenon occurs when the electrostatic force has become equal to the surface tension of the liquid and at that point the droplet changes shape to an elliptically shaped Taylor cone that is drawn out toward the mass spectrometer orifice along the axis of the fluid flow. The gap distance between the spray tip and MS orifice, as well as the alignment of the spray tip is therefore critical, and so the gap should be kept between 3 and 6 mm. The initial fine droplets sprayed from the tip shrink by solvent evaporation, and ions, which are involatile, are retained in the shrinking droplet. The increase in the repulsive forces between the excess charges in the droplet eventually overcome cohesive forces in the droplet to cause disintegration into smaller droplets or columbic explosion (22). A small orifice in the counter electrode then allows some of the ions from solution to enter the vacuum chamber of the mass spectrometer for mass analysis. 3. In cases where the electrophoresis buffer is incompatible with electrospray, a sheath liquid or make-up solution can be introduced through a side arm microchannel as shown in Fig. 3 (see Note 13). The sample meets the make-up solution flow from a syringe pump at the intersection of the channel from D and the electrophoresis main channel. A 75-µm I.D and 360-µm O.D fused silica capillary column is connected at one end to reservoir D on the microchip through one of the Upchurch nanoport assemblies, and the other end to the mass spectrometer syringe pump. A 250-µL syringe is filled with the make-up or sheath solution, and delivered on to the chip at a rate lower than the EOF stream, which is typically less than 100 nL/min. A flow splitter is connected between the microchip and the syringe pump to achieve the required low flow rates. If the flow rate of the make-up solution becomes too high, the solution can migrate backwards into the separation channel toward the separation buffer reservoir and the separated sample components never reach the spray tip. The flow rate should be kept between 50 and 100 µL/min for electrospray conditions described in this chapter. If the separation buffer is ESI compatible, then the use

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of this make-up solution can be eliminated (see Note 14). The make-up solution is used to modify the electrophoresis buffer to meet the requirements of electrospray conditions. In cases where the electrophoresis buffer is too concentrated (which is usually the case because electrophoresis buffer needs to be highly conductive for significant EOF), the make-up solution is used to dilute the buffer for the ESI-MS, which requires a lower concentration of buffer to minimize background noise. 4. The EOF, together with the hydrodynamic flow stream of the make-up solution, deliver the sample to the spray tip of the fused silica capillary emitter where electrospray occurs. The CE voltage applied at reservoir A can be decoupled from the electrospray process by applying the ESI voltage at reservoir D (see Note 15). 5. Separated sample components are detected using positive ion mode MS because separation is at low pH and the sample ions are predominantly positive ions.

4. Notes 1. In photolithography, alternative borosilicate glass types such as D263, Corning 0211, or Schott Borofloat can be used. 2. For smaller channel features, more UV exposure time is required. Thus, UV irradiation time is dependant upon the time needed to create preferable dimensions of the smallest width of the channels. 3. Etching time varies with types of glass and the desired etching depth. Soda-lime glass etches faster than Corning 0211, D263, and the Schott Borofloat. 4. It should be realized that the anisotropic etching of the glass substrates by HF normally produces trapezoidal-shaped channels. The stylus instrument is used to measure the width at half the height and depth of the channels. 5. In the event that the bonding process yields interfering Newton rings or a weak bond, place the chip in water for several hours, depending on the strength of the bond that would have been formed, and insert a wedge between the glass plates to separate the two. Perform the washing and drying cycle again in the wafer washer, after which bonding steps 6–7 (Subheading 3.1.3.) are repeated. If there are a few Newton rings that do not cross the channel network, proceed on to the hightemperature bonding of the microchip. 6. When drilling the vertical access holes do not use any liquid to collect glass chippings or aid the drilling process, instead simply fan accumulating particulates away. 7. Water is filled in all the channels before the drilling of horizontal dead volume connection because the movement of water denotes the success of a connection and the water prevents the glass particles from the drilling process from getting lodged in the channel during the course of the drilling process. 8. Ensure that the capillary tip end is flattened by the use of a sandpaper or emery cloth to avoid use of jagged edge capillaries. View the tip with a microscope to verify the flatness.

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9. A variety of coatings can be used to modify the surface of the microchip to enhance or suppress EOF and to prevent protein adsorption (23). Examples are ([acryloylamino]propyl)trimethylammonium chloride (BCQ) and (3-aminopropyl) silane, which have been used to impart a positive charge on the surface of the microchip channels for EOF enhancement through the microchannel at low pH (24,25). Linear polyacrylamide and polyvinyl alcohol (PVA) have also been used for EOF suppression and minimization of sample adsorption to channel surface for protein and peptide separations (26). 10. The length of the spray capillary should be kept at a minimum because of the electrokinetically induced pressures that develop at the microchip–capillary junction. The pressure term introduced by the capillary reduces the EOF through the tip (27). 11. Sample leakage into the separation channel can be avoided by applying small biasing voltages to reservoirs B and C. 12. The injection plug length can be varied by increasing the distance between the side channels of the double-tee injector in the fabrication process. The pinched injection strategy can also be used for injection using a cross or single-tee instead of a double-tee (28). Continuous infusion of the sample into the MS can be achieved by applying a negative voltage to the sample reservoir (B) and leaving A and C floating. 13. Although a number of researchers have made use of a side arm microchannel to introduce make-up solution (12,13,29), other groups have used on-chip porous microjunctions (27). Make-up solutions can also be introduced through external liquid junctions at the outlet of the CE microchannel. 14. There are a few reports in literature of direct coupling of microchip CE and ESIMS with direct generation of electrospray from EOF through the electrophoresis channel, without the use of a make-up solution. An example is a report by Gobry et al., where a polymer-based microchip was directly coupled to the MS for detection of small ions, but the performance of the interface was hampered by the large dead volume from the droplet that formed at the outlet of the microchannel (30). Lazar et al. also demonstrated electrospray from a glass microchip that made use of extremely low EOF rates (20–30 nL/min) to achieve subattomole sensitivity for proteins and peptides. The device consisted of a nanospray tip perpendicularly attached to the microchip surface (31). However, most interfaces in literature, even those that use an attached capillary, normally make use of make-up solution to modify the electrophoresis buffer to achieve the best possible conditions for stable electrospray. 15. The separation voltage can be decoupled from the high voltage used for electrospray. Other than the method described in this chapter, the ESI voltage can be applied and controlled at the spray tip using either a direct electrical connection through a gold-coated capillary used by Harrison and co-workers (12,13,29,32,33), a liquid junction used by Karger et al. (11,26,34,35), or a sheath flow junction (12). The gold-coated capillary and liquid junction configurations are shown in Fig. 4.

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Fig. 4. (A) Schematic diagram of a microchip-capillary electrophoresis mass spectrometry configuration using a gold-coated spray tip. The electrospray ionization (ESI) voltage is applied via an electrode attached to the gold coating. (B) A schematic representation of a liquid junction. In this case the ESI voltage and the make-up solution is applied at the liquid junction.

References 1. Waters, L. C., Jacobson, S. C., Kroutchinina, N., Khandurina, J., Foote, R. S., and Ramsey, J. M. (1998) Microchip device for cell lysis, multiplex PCR amplification, and electrophoretic sizing. Anal. Chem. 70, 158–162. 2. Manz, A., Harrison, D. J., Verpoorte, E. M. J., et al. (1992) Planar chips technology for miniaturization and integration of separation techniques into monitoring systems. Capillary electrophoresis on a chip. J. Chromatogr. 593, 253–258. 3. Ross, D. and Locascio, L. E. (2002) Microfluidic temperature gradient focusing. Anal. Chem. 74, 2556–2564.

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4. Wang, C., Oleschuk, R., Ouchen, F., Li, J., Thibault, P., and Harrison, D. J. (2000) Integration of immobilized trypsin bead beds for protein digestion within a microfluidic chip incorporating capillary electrophoresis separations and an electrospray mass spectrometry interface. Rapid Commun. Mass Spectrom. 14, 1377–1383. 5. Manz, A. and Becker, H. (1997) Parallel capillaries for high throughput in electrophoretic separations and electroosmotic drug discovery systems. Transducers 97, International Conference on Solid-State Sensors and Actuators, Chicago, June 16–19, 2, 915–918. 6. Liu, H., Felten, C., Xue, Q., et al. (2000) Development of multichannel devices with an array of electrospray tips for high-throughput mass spectrometry. Anal. Chem. 72, 3303–3310. 7. Tang, K., Lin, Y., Matson, D. W., Kim, T., and Smith, R. D. (2001) Generation of multiple electrosprays using microfabricated emitter arrays for improved mass spectrometric sensitivity. Anal. Chem. 73, 1658–1663. 8. Gelpi, E. (2002) Interfaces for coupled liquid-phase separation/mass spectrometry techniques. An update on recent developments. J. Mass. Spectrom. 37, 241–253. 9. Xue, Q., Foret, F., Dunayevskiy, Y. M., Zavracky, P. M., McGruer, N. E., and Karger, B. L. (1997) Multichannel microchip electrospray mass spectrometry. Anal. Chem. 69, 426–430. 10. Ramsey, R. S. and Ramsey, J. M. (1997) Generating electrospray from microchip devices using electroosmotic pumping. Anal. Chem. 69, 1174–1178. 11. Zhang, B., Liu, H., Karger, B. L., and Foret, F. (1999) Microfabricated devices for capillary electrophoresis-electrospray mass spectrometry. Anal. Chem. 71, 3258–3264. 12. Li, J., Thibault, P., Bings, N. H., 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. Anal. Chem. 71, 3036–3045. 13. Li, J., Kelly, J. F., Chernushevich, I., Harrison, D. J., and Thibault, P. (2000) Separation and identification of peptides from gel-isolated membrane proteins using a microfabricated device for combined capillary electrophoresis/nanoelectrospray mass spectrometry. Anal. Chem. 72, 599–609. 14. Figeys, D., Ning, Y., and Aebersold, R. (1997) A microfabricated device for rapid protein identification by microelectrospray ion trap mass spectrometry. Anal. Chem. 69, 3153–3160. 15. Schultz, G. A., Corso, T. N., Prosser, S. J., and Zhang, S. (2000) A fully integrated monolithic microchip electrospray device for mass spectrometry. Anal. Chem. 72, 4058–4063. 16. Wen, J., Lin, Y., Xiang, F., Matson, D. W., Udseth, H. R., and Smith, R. D. (2000) Microfabricated isoelectric focusing device for direct electrospray ionizationmass spectrometry. Electrophoresis 21, 191–197. 17. Wilm, M. and Mann, M. (1996) Analytical properties of the nanoelectrospray ion source. Anal. Chem. 68, 1–8.

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18. Valaskovic, G. A., Kelleher, N. L., Little, D. P., Aaserud, D. J., and McLafferty, F. W. (1995) Attomole-sensitivity electrospray source for large-molecule mass spectrometry. Anal. Chem. 67, 3802–3805. 19. Jacobson, S. C., Hergenroder, R., Koutny, L. B., Warmack, R. J., and Ramsey, J. M. (1994) Effects of injection schemes and column geometry on the performance of microchip electrophoresis devices. Anal. Chem. 66, 1107–1113. 20. Khaledi, M. G. (1998) High-Performance Capillary Electrophoresis: Theory, Techniques, and Applications, Wiley, New York, NY. 21. Chiem, N., Lockyear-Shultz, L., Andersson, P., Skinner, C., and Harrison, D. J. (2000) Room temperature bonding of micromachined glass devices for capillary electrophoresis. Sens. Actuators B Chem. B63, 147–152. 22. Gaskell, S. J., Bolgar, M. S., Riba, I., and Summerfield, S. G. (1997) Electrospray ionization: theory and application. NATO Adv. Stud. Inst. Ser C: Mathematical and Physical Sciences 504, 3–16. 23. Belder, D. and Ludwig, M. (2003) Surface modification in microchip electrophoresis. Electrophoresis 24, 3595–3606. 24. Li, J., LeRiche, T., Tremblay, T. -L., et al. (2002) Application of microfluidic devices to proteomics research: identification of trace-level protein digests and affinity capture of target peptides. Mol. Cell Proteomics 1, 157–168. 25. Figeys, D. and Aebersold, R. (1998) Nanoflow solvent gradient delivery from a microfabricated device for protein identifications by electrospray ionization mass spectrometry. Anal. Chem. 70, 3721–3727. 26. Zhang, B., Foret, F., and Karger, B. L. (2000) A microdevice with integrated liquid junction for facile peptide and protein analysis by capillary electrophoresis/electrospray mass spectrometry. Anal. Chem. 72, 1015–1022. 27. Lazar, I. M., Ramsey, R. S., Jacobson, S. C., Foote, R. S., and Ramsey, J. M. (2000) Novel microfabricated device for electrokinetically induced pressure flow and electrospray ionization mass spectrometry. J. Chromatogr. 892, 195–201. 28. Alarie, J. P., Jacobson, S. C., and Ramsey, J. M. (2001) Electrophoretic injection bias in a microchip valving scheme. Electrophoresis 22, 312–317. 29. Li, J., Wang, C., Kelly, J. F., Harrison, D. J., and Thibault, P. (2000) Rapid and sensitive separation of trace level protein digests using microfabricated devices coupled to a quadrupole—time-of-flight mass spectrometer. Electrophoresis 21, 198–210. 30. Gobry, V., Van Oostrum, J., Martinelli, M., et al. (2002) Microfabricated polymer injector for direct mass spectrometry coupling. Proteomics 2, 405–412. 31. Lazar, I. M., Ramsey, R. S., Sundberg, S., and Ramsey, J. M. (1999) Subattomolesensitivity microchip nanoelectrospray source with time-of-flight mass spectrometry detection. Anal. Chem. 71, 3627–3631. 32. Deng, Y., Henion, J., Li, J., Thibault, P., Wang, C., and Harrison, D. J. (2001) Chip-based capillary electrophoresis/mass spectrometry determination of carnitines in human urine. Anal. Chem. 73, 639–646. 33. Li, J., Tremblay, T. -L., Thibault, P., Wang, C., Attiya, S., and Harrison, D. J. (2001) Integrated system for high-throughput protein identification using a micro-

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fabricated device coupled to capillary electrophoresis/nanoelectrospray mass spectrometry. Eur. J. Mass. Spectrom. 7, 143–155. 34. Zhang, B., Foret, F., and Karger, B. L. (2001) High-throughput microfabricated CE/ESI-MS: Automated sampling from a microwell plate. Anal. Chem. 73, 2675–2681. 35. Foret, F., Zhou, H., Gangl, E., and Karger, B. L. (2000) Subatmospheric electrospray interface for coupling of microcolumn separations with mass spectrometry. Electrophoresis 21, 1363–1371.

8 Interfacing Amperometric Detection With Microchip Capillary Electrophoresis R. Scott Martin Summary Amperometric detection is a sensitive and selective way to monitor separations in microchip capillary electrophoresis (CE). This review contains 78 references and will educate the reader of the issues that are involved with interfacing amperometric detection and microchip CE. These issues include special injection protocols, separation mechanisms, and ways to integrate the working electrode with the separation channel. Some useful biological applications of the technique will also be described. Key Words: Amperometric; electrochemistry; capillary electrophoresis; microchip; micrototal analysis system; portable analysis system; microfluidics.

1. Introduction Capillary electrophoresis (CE) in the microchip format has come a long way over the past decade. Initial demonstrations of the technique (1–4) showed the numerous benefits of the microchip format that include fast analysis times, the use of high separation field strengths, minute consumption of solvents, and the possibilities for disposable/portable devices. These numerous advantages have lead to many exciting applications of microchip CE including fully integrated multichannel separation-based immunoassays (5), complex two-dimensional separations (6), and high-throughput 384-channel DNA separations (7). The small volume ( 3d the tanh function is approx 1 and can be dropped. For channels where w > 10d the entire last term in the equation is approx 1. This equation can be used to design the appropriate flow resistances into the channel manifold on the microfluidic device.

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3.2. M icrochip Fabrication

3.2.1. Glass Etching (28–30) 1. Photomasks with the appropriate channel designs can be drawn using AutoCAD LT2000 or another CAD drawing program and saved as .dwg files. Eight 2 × 1-in. channel designs are generally drawn for each 4 × 4 in. mask. The lines on the mask should be 10–20 µm wide. Glass etches isotropically, so when etching channels made from masks with a line width of lm, the width of the channels will be approx 2d + lm wide at the top and lm wide at the bottom where d is the depth of the channel. The .dwg files containing the mask designs can be electronically sent to the variety of photomask manufacturers including HTA Photomask (San Jose, CA, www.htaphotomask.com). 2. Align the photomask with a photomask blank on a flood exposure system with the chrome-plated side touching the resist of the blank substrate and expose for 5 s using a flood exposure system (see Note 8). The exposure time may vary with the photoresist used and also with the age of the lamp. 3. Develop the exposed masks in Microposit 453 for 60 s, rinse with ultrapure (18 MΩ) water for 1 min, and dry under argon or some other dry, inert gas. 4. Etch the exposed chrome on the masks using the chrome etchant for 3–4 min, and then rinse with ultrapure water. 5. Remove undeveloped resist with acetone. Rinse with water and dry using argon. Repeat resist removal procedure if necessary. 6. Place the substrate in a stirred 1:10 BOE solution in an appropriate plastic container. The etch rate is very sensitive to temperature and will change with the age of the solution. Average etch rates are approx 0.25–0.50 µm/min. Remove the substrate periodically from the solution (see Note 9), rinse with water, blow dry with argon, and measure the depth of the channels with a stylus-based profiler (TencorKLA Alphastep IQ or similar instrument). When the channels have been etched to a depth of approx 18 µm, remove the substrate, rinse with water and then 95% ethanol, and dry (see Note 10). 7. Remove the chrome on the substrate by agitating in chrome etchant for approx 10 min. Rinse thoroughly with water. Blow dry. 8. Cut substrate into eight 2 × 1-in. chips using a glass dicing saw. Also cut a blank substrate (i.e., uncoated clean glass) into 2 × 1-in. rectangles for use as the cover plates. On the cover plate, drill access holes to the channels using diamond-coated drill bits or an ultrasonic mill.

3.2.2. Bonding (28–30) 1. Sonicate substrate and cover plate in a detergent solution for 10–20 min, rinse with ultrapure water, and dry. 2. Sonicate in acetone for 10 min and then blow dry. 3. Etch in 1:10 BOE solution for 1 min, rinse with water, and place directly into the heated hydrolysis solution. 4. The hydrolysis solution should be heated to approx 50°C prior to the immersion of the substrates, and the substrates should soak in this solution for 20 min (see Notes 11 and 12).

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5. Remove the substrates from the hydrolysis solution and sonicate in running ultrapure water for 15 min. 6. Remove the substrate and glass while the water is still running and join together. This step should be performed in a laminar flow hood or cleanroom. 7. Let sit at room temperature for 5 min and then place in a 95°C oven for 30 min. After 30 min remove from oven to see if bonding has occurred. If bonding looks good then replace in oven for another 6 h. Nonbonded chips easily come apart. Partially bonded chips usually show a series of Newton’s rings (diffraction patterns) in the areas on the chip where bonding has failed. 8. To permanently anneal the chips, place them in an oven and raise the temperature to the annealing temperature for that type of glass. For soda-lime glasses that temperature is generally 525–550°C, for borofloat glasses it is 625–675°C, and for quartz it is approx 1100°C. The following procedure can be used for soda-lime glasses. Raise the temperature of the oven from 95 to 200°C at a rate of 0.2–0.5°C/min. From 200 to 525°C the temperature can be ramped at 5–10°C/min (see Note 13). Let sit at 525°C for 6 h and then passively cool to room temperature (see Note 14). 9. Upchurch Nanoport® reservoirs (Upchurch Scientific, Oak Harbor, WA) can be added to the chip to increase the fluid volume of the reservoir. These reservoirs also provide a handy way to couple the waste channel to the syringe pump.

3.3. Fluorescence D etection Setup

Several fluorescent detection schemes have be reported in the literature (29,31–33). Below is an outline for setting up a basic single point detection system. The basic support structure for an epi-illumination setup (Fig. 2) can be built from equipment obtained from Thorlabs (Newton, NJ; www.thorlabs.com) and Linos Photonics (Milford, MA; www.linos-photonics.com). For Calcein, Oregon green, and carboxyfluorescein, the 488-nm line on an argon ion laser is used to excite the fluorescence. The laser beam is reflected off of a dichroic mirror and focused through an inexpensive ×40 ELWD microscope objective. The same objective is used to collect the fluorescent emission from the dyes. The fluorescent emission is passed through the dichroic and imaged onto a spatial filter (1-mm pinhole). The spatially filtered emission is then passed through a 530-nm bandpass filter and finally detected at a PMT. A 488-nm notch filter can also be placed into the light path for better noise rejection. The signal from the PMT is amplified by the low noise current amplifier and digitized by the multifunction I/O card. To detect the intact cells just prior to lysis, in addition to the lysate in the separation channel, a beam splitter is used to split the light from the laser. The split light is then focused into the microchip channel approx 10–20 µm upstream of the lysis/injection cross. The fluorescence from the intact cells as they pass through the focused laser beam is collected by a second microscope objective, passed through a spatial and spectral filter, and detected by a PMT.

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Fig. 2. Single-point detection schematic (see Subheading 3.3. for details).

3.4. Channel Wall Coating

To prevent cell and cell lysate adhesion to the channel walls, the following channel-coating procedure is used. This procedure also significantly reduces the electroosmotic flow in the separation channel (see Note 15). 1. Prepare the channels in the chip to be coated by flushing 1 M NaOH through the channels for 5 min. Follow this by flushing water through the channels for 5 min. Dry the channels out by pulling air through them. Finally put the chip in a 110°C oven for 2 h. 2. Mix 1 g of Sylgard 184 part A with 0.1 g of Sylgard 184 part B. 3. Dilute this mixture to 20% (v/v) in hexane. 4. Place mixture in all of the reservoirs and apply a vacuum to the waste reservoir for 5 min.

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5. Replace the diluted Sylgard mixture with neat hexane and aspirate through the chip for 10 min to remove excess Sylgard mixture. 6. Dry the chip. 7. Place in a 65°C oven overnight to cure (see Note 16). 8. Just prior to filling the chip with the separation buffer for the cell analysis, aspirate a solution of 30% (v/v) Pluronic F-127 in water through the chip for 20 min.

3.5. Cell Culture (34) 1. A supplemented RMPI 1640 medium is used to culture the Jurkat cells as reported in the Subheading 2.1. 2. An inoculate of cells is added to 10 mL of the supplemented media in 25-mL polystyrene culture flasks. 3. The cultures should be maintained at 37°C with a CO2 concentration of 5%. 4. The cells should be passed when the cell density reaches 106/mL. Cell density can be determined using a hemacytometer (34). 5. Cells should not be used until after the third passage.

3.6. Cell Loading 1. Dyes: Oregon green 488 carboxylic acid diacetate (OGCA-D), carboxyfluorescein diacetate (CF-D), and Calcein AM (C-AM) are all fluorogenic and cell membrane permeable. 2. Take a 1-mL aliquot of cells in media and spin the cells down in a centrifuge. 3. Decant the media and replace with extracellular buffer (see Subheading 2.1.) at room temperature. 4. Gently resuspend the cells at a concentration of about 1 × 106/mL. 5. OGCA-D should be constituted at a concentration of 20 µM in the extracellular buffer containing the cells and incubated at room temperature for 10 min. CF-D and C-AM should be constituted at a concentration of 1 µM in the extracellular buffer containing the cells and incubated at room temperature for 10 min (12,15). At these concentrations most of the dye diffuses into the cell and there is no need to replace the buffer. This step can be performed in the cell reservoir on the chip (see Subheading 3.7., step 2).

3.7. Cell Transport and Lysis 1. Fill all of the reservoirs with the separation buffer and flush through the chip to fill all of the channels. 2. Remove separation buffer from the cell reservoir and add either the loaded or unloaded cells in extracellular buffer. If the cells are unloaded, then add a sufficient amount of dye to bring the final dye concentration in the reservoir to that specified in Subheading 3.6., step 5. Unloaded cells are spun down in a centrifuge and resuspended in extracellular buffer at a concentration of approx 1 × 106/mL (see Note 17). 3. Generate a subambient pressure at the cell waste reservoir using a syringe pump.

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Fig. 3. Dual point detection example. The bottom trace is from the single point detection system 3 mm down the separation channel from the lysis intersection. The peaks resulting from each individual cell are underlined and numbered. The top trace is the fluorescent signal from the intact cells just prior to lysis. The arrows from the top trace point to the peak envelopes in the bottom trace to which they correspond. The trace has been inverted to more easily see the correspondence between the signals. (Reprinted with permission from ref. 15 © 2003 American Chemical Society.) 4. Increase the subambient pressure until the cells are flowing at a rate of approx 1 mm/s. To measure the cell velocities, they can be imaged as discussed in Subheading 3.8. 5. Apply an electric field across the separation channel using the output from a waveform generator amplified by an AC amplifier (see Subheading 2.2. and Note 18). The electric potential should be applied as a square wave at a frequency of approx 75 Hz. The field strength should vary from approx 450 to 900 V/cm at the nadir and apex of the wave. The time-dependent electric field is used to reduce Joule heating problems. 6. With the first part of the single point detection system located 3–5 mm downstream of the lysis intersection in the separation channel and the second part just above the lysis intersection, a successful experiment results in data which can be seen in Fig. 3 (see Subheading 3.3. and Note 19). The separation should be completed in less than 5 s with migration time reproducibilities for all peaks of less than 1% (15).

3.8. Chip and Cell Imaging

To ascertain if the cells are moving properly through the chip and to optimize the lysis and injection efficiencies, the cross intersection of the microfluidic

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device can be imaged. This is performed most conveniently using an inverted microscope equipped with an extra long distance condenser lens and an epifluorescence attachment. Such a setup allows plenty of room above the microscope stage for the microchip and its accessory components. The microchip is simply taped to the glass stage ring and imaged. 3.9. Lysate Injection Efficiency M easurements

Lysate injection efficiency can be quantitated by looking for fluorescence in the waste channel when the electric field is on. When the electric field is off, one should be able to detect the fluorescence from intact cells migrating down the waste channel; however, once the voltage has been applied to the chip, then the fluorescence should disappear. If some fluorescence is seen, then the cell flow rate needs to be decreased to allow more time for the lysate to migrate into the injection channel once the cell is lysed. 3.10. Channel Wall Coating D urability

The channel wall coating has a limited lifetime. When the analyte peak migration times and peak shapes begin to degrade, then the chip coating needs to be replaced. The coating may last several days before needing to be replaced. To recoat a chip, the old coating is first pyrolyzed at 500°C in an oven for 8 h. The channel is then rinsed with water and dried. The coating procedure in Subheading 3.4. is then repeated. 3.11. D ata Collection and Analysis

Simple data collection and analysis routines can be written using LabVIEW (National Instruments, Austin, TX) and Igor Pro (Wavemetrics, Lake Oswego, OR). 4. N otes 1. Many of the materials used for the experiments described below entail the use of potentially hazardous substances, so care should be taken in their use. One should be familiar with the Material Safety Data Sheet sheets of each component, use the proper personal protective gear, and perform the experiments in a fume hoods where necessary. 2. The photoresist developer is a strong base, so care should be exercised when using this solution along with wearing the proper personal protection. 3. The glass etching solution contains strong acids, so care should be exercised when using this solution along with wearing the proper personal protection. 4. This solution contains hydrofluoric acid, so extreme care should be exercised when using this solution along with wearing the proper personal protection. 5. This solution contains both a strong base and oxidizer, so care should be exercised when using this solution along with wearing the proper personal protection. The

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7. 8. 9. 10. 11.

12. 13. 14. 15. 16. 17.

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19.

Culbertson exothermic nature of the hydrolysis reaction can further raise the temperature upon immersion of the glass slides. The optiprep makes the extracellular buffer isopycnic so that the cells do not settle out in the cell reservoir over time. This helps to keep that rate of cell entry into the main chip channel constant over the course of the entire experiment. The Jurkat cells used for these experiments are very sensitive to electric fields and, therefore, cannot be transported via electrokinesis. The chrome on the photomask is only on the order of 50- to 100-nm thick, so care should be exercised in handling the masks to prevent scratching. Place substrate in a plastic container to prevent dripping of hydrofluoric acid. If etch rates are uneven from one edge of the substrate to another, the substrate can be rotated by 90° at periodic intervals. The hydrolysis of the surface of the glass slide is exothermic. Depending on the volume of the hydrolysis solution and the number of substrates immersed, a considerable temperature rise in the solution can be seen. The hydrogen peroxide in the solution has a limited lifetime of only a few hours, so the solution needs to be regularly replaced. The annealing temperature varies slightly with different types of glasses, so the final temperature may need to be adjusted slightly. A weight may also be placed on the chip to help assure good bonding. An alternative procedure developed by Harrison’s group for pyrolyzing poly(dimethylsiloxane) on the channel wall surface works equally well (35). The curing should be performed in an explosion proof oven. The cells are viable in the extracellular buffer at room temperature for several hours; however, the dyes do begin to bleed out of the cells approx 1 h after loading. Experiments, therefore, should be carried out within 1 h of the loading process. The high voltages used to lyse the cells and then to separate the lysate are potentially dangerous. For this reason it is prudent to include an interlock system in the experimental setup to prevent operators from accidentally shocking themselves during an experiment. Cells enter the main channel of the microchip at random time intervals; so the average distance between cells in the main channel and the average time between lysis events also varies randomly. For the analytes released from one cell to be completely separated from the cell prior to it and after it, a certain average distance between cells must be maintained. For detection distances of 3–5 mm in the system described here, the optimal cell concentration was 1 × 106 cells/mL.

Acknowledgments

This work was supported by the National Institutes of Health by Grant RO1GM067905. References 1. Lodish, H., Baltimore, D., Berk, A., Zipursky, S. L., Matsudaira, P., and Darnell, J. (1995) Molecular Cell Biology, 3rd Edition, Scientific American Books, New York, NY.

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2. Zabzbyr, J. L. and Lillard, S. J. (2001) New Approaches to Single-cell Analysis by Capillary Electrophoresis. Trends Analyt. Chem. 20, 467–476. 3. Sims, C. E., Meredith, G. D., Krasieva, T. B., Berns, M. W., Tromberg, B. J., and Allbritton, N. L. (1998) Laser-micropipet combination for single-cell analysis. Anal. Chem. 70, 4570–4577. 4. Berridge, M. J. (1993) Inositol trisphosphate and calcium signalling. Nature 361, 312–325. 5. Chen, G. and Ewing, A. G. (1997) Chemical analysis of single cells and exocytosis. Crit. Rev. Neurobiol. 11, 59–90. 6. Hsieh, S., Dreisewerd, K., van der Schors, R. C., et al. (1998) Separation and identification of peptides in single neurons by microcolumn liquid chromatographymatrix-assisted laser desorption/ionization time-of-flight mass spectrometry and postsource decay analysis. Anal. Chem. 70, 1847–1852. 7. Hsieh, S. and Jorgenson, J. W. (1997) Determination of enzyme activity in single bovine adrenal medullary cells by separation of isotopically labeled catecholamines. Anal. Chem. 69, 3907–3914. 8. Pihel, K., Hsieh, S., Jorgenson, J. W., and Wightman, R. M. (1995) Electrochemical detection of histamine and 5-hydroxytryptamine at isolated mast cells. Anal. Chem. 67, 4514–4521. 9. Swanek, F. D., Ferris, S. S., and Ewing, A. G. (1997) Capillary Electrophoresis for the Analysis of Single Cells: Electrochemical, Mass Spectrometric, and Radiochemical Detection. In: Handbook of Capillary Electrophoresis, (Khaledi, M. G., ed.), CRC Press, Inc., Boca Raton, FL, pp. 495–521. 10. Ewing, A. G., Chen, T. -K., and Chen, G. (1995) Voltammetric and Amperometric Probes for Single-Cell Analysis. In: Voltammetric Methods in Brain Systems, (Boulton, A., Baker, G., and Adams, R. N., eds.), Humana Press, Totowa, NJ, pp. 269–304. 11. Lillard, S. J. and Yeung, E. S. (1997) Capillary Electrophoresis for the Analysis of Single Cells: Laser-Induced Fluorescence Detection. In: Handbook of Capillary Electrophoresis (Khaledi, M. G., ed.), CRC Press, Inc., Boca Raton, FL, pp. 523–544. 12. Han, F., Wang, Y., Sims, C. E., et al. (2003) Fast electrical lysis of cells for capillary electrophoresis. Anal. Chem. 75, 3688–3696. 13. Meredith, G. D., Sims, C. E., Soughayer, J. S., and Allbritton, N. L. (2000) Measurement of kinase activation in single mammalian cells. Nature Biotech. 18, 309–312. 14. Lee, C. L., Linton, J., Soughayer, J. S., Sims, C. E., and Allbritton, N. L. (1999) Localized measurement of kinase activation in oocytes of Xenopus laevis. Nat. Biotechnol. 17, 759–762. 15. McClain, M. A., Culbertson, C. T., Jacobson, S. C., Allbritton, N. L., Sims, C. E., and Ramsey, J. M. (2003) Microfluidic devices for the high-throughput chemical analysis of cells. Anal. Chem. 75, 5646–5655. 16. Fu, A. Y., Chou, H. -P., Spence, C., Arnold, F. H., and Quake, S. R. (2002) An integrated microfabricated cell sorter. Anal. Chem. 74, 2451–2457. 17. Fu, A. Y., Spence, C., Scherer, A., Arnold, F. H., and Quake, S. R. (1999) A microfabricated fluorescence-activated cell sorter. Nature Biotech. 17, 1109–1111.

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18. McClain, M. A., Culbertson, C. T., Jacobson, S. C., and Ramsey, J. M. (2001) Flow cytometry of Escherichia coli on microfluidic devices. Anal. Chem. 73, 5334–5338. 19. Roper, M. G, Shackman, J. G., Dahlgren, G. M., and Kennedy, R. T. (2003) Microfluidic chip for continuous monitoring of hormone secretion from live cells using an electrophoresis-based immunoassay. Anal. Chem. 75, 4711–4717. 20. Wheeler, A. R., Throndset, W. R., Whelan, R. J., et al. (2003) Microfluidic device for single-cell analysis. Anal. Chem. 75, 3581–3586. 21. Fuhr, G. R. and Reichle, C. (2000) Living cells in opto-electrical cages. Trends Analyt. Chem. 19, 402–409. 22. Yang, J., Huang, Y., Wang, X. -B., Becker, F. F., and Gascoyne, P. R. C. (1999) Cell separation on microfabricated electrodes using dielectrophoretic/gravitational field-flow fractionation. Anal. Chem. 71, 911–918. 23. Yang, M., Li, C. -W., and Yang, J. (2002) Cell docking and on-chip monitoring of cellular reactions with a controlled concentration gradient on a microfluidic device. Anal. Chem. 74, 3991–4001. 24. Schilling, E. A., Kamholz, A. E., and Yager, P. (2002) Cell lysis and protein extraction in a microfluidic device with detection by a fluorogenic enzyme assay. Anal. Chem. 74, 1798–1804. 25. Muller, T., Gradl, G., Howitz, S., Shirley, S., Schnelle, T., and Fuhr, G. (1999) A 3-D microelectrode system for handling and caging single cells and particles. Biosens. Bioelectron. 14, 247–256. 26. Li, P. C. H. and Harrison, D. J. (1997) Transport, manipulation, and reaction of biological cells on-chip using electrokinetic effects. Anal. Chem. 69, 1564–1568. 27. White, F. M. (1991) Viscous Fluid Flow, Second Edition, McGraw-Hill, New York, NY. 28. Fortina, P., Cheng, J., Kricka, L. J., et al. (2001) DOP-PCR amplification of whole genomic DNA and microchip-based capillary electrophoresis. In: Capillary Electrophoresis of Nucleic Acids, Volume 2, (Mitchelson, K. R. and Cheng, J., eds.), Humana Press, Totowa, NJ, pp. 211–219. 29. Jacobson, S. C., Hergenröder, R., Koutny, L. B., Warmack, R. J., and Ramsey, J. M. (1994) Effects of injection schemes and column geometry on the performance of microchip electrophoresis devices. Anal. Chem. 66, 1107–1113. 30. Stjernstrom, M. and Roeraade, J. (1998) Method for fabrication of icrofluidic systems in glass. J. Micromech. Microeng. 8, 33–38. 31. Mets, U. and Rigler, R. (1994) Submillisecond detection of single rhodamine molecules in water. J. Fluoresc. 4, 259–264. 32. Nie, S., Chiu, D. T., and Zare, R. N. (1994) Probing individual molecules with confocal fluorescence microscopy. Science 266, 1018–1021. 33. Schrum, D. P., Culbertson, C. T., Jacobson, S. C., and Ramsey, J. M. (1999) Microchip flow cytometry using electrokinetic focusing. Anal. Chem. 71, 4173–4177. 34. McAteer, J. A. and Davis, J. (1994) Basic cell culture technique and the maintenance of cell lines. In: Basic Cell Culture (Davis, J., ed.), Oxford University Press, New York, NY, pp. 93–148. 35. Badal, M. Y., Wong, M., Chiem, N., Salimi-Moosavi, H., Harrison, D. J. (2000) J. Chromatogr. A 947, 277–286.

15 Rapid DNA Amplification in Glass Microdevices Christopher J. Easley, Lindsay A. Legendre, James P. Landers, and Jerome P. Ferrance Summary The polymerase chain reaction (PCR) for amplification of DNA has become a very useful tool in scientific research and analytical laboratories, yet conventional techniques are time-consuming, and the reagents are expensive. Miniaturization of this technique has the potential to drastically reduce amplification time and reagent consumption while simultaneously improving the efficiency of the reaction. Increasing the surface area-to-volume ratio using microfluidic reaction chambers allows homogeneous solution temperatures to be achieved much more rapidly than in conventional heating blocks. Employing infrared radiation to selectively heat the reaction solution can additionally reduce the time and energy needed for thermocycling; the reaction container is not heated and can even serve as a heat sink for enhancement of cooling. Microchip systems also provide the potential for fabrication of structures for additional processing steps directly in line with the PCR chamber. Not only can amplification be integrated with product separation and analysis, but sample preparation steps can also be incorporated prior to amplification. The ultimate goal is a miniature total-analysis-system with seamlessly coupled sample-in/answer-out capabilities that consumes very low volumes of reagents and drastically reduces the time for analysis. This chapter will focus on the materials and methods involved in simple straight-channel microchip PCR on glass substrates using non-contact thermocycling. Key Words: PCR; microchip; infrared; non-contact.

1. Introduction Since its first reported use in 1985 (1), and particularly since the advent of thermally stable polymerase enzymes, the polymerase chain reaction (PCR) has rapidly become a basic and essential tool in biochemical research and analytical laboratories. PCR has developed into an invaluable technique in clinical diagnostics for detecting infectious agents, and is being applied for detection of genetic changes associated with specific diseases. The reaction has also been widely developed in the forensic community for identification of suspects or victims in criminal investigations and can be used for identification of victims From: Methods in Molecular Biology, vol. 339: Microchip Capillary Electrophoresis: Methods and Protocols Edited by: C. S. Henry © Humana Press Inc., Totowa, NJ

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of terrorist attacks. More recent motivation for the application of PCR has been found in the rapid detection of agents of bioterrorism for accurate and focused response to attacks. PCR is an in vitro technique allowing amplification of a specific DNA fragment of known length. Short oligonucleotides, called primers, which are complementary to the 3′ sequences at the ends of the specific DNA fragment of interest, are mixed with nucleotides, a small amount of template DNA, and Taq DNA polymerase enzyme in the appropriate buffer. The temperature of the reaction mixture is then cycled to denature the double stranded DNA, allow annealing of primers at the ends of the DNA sequence to be amplified (typically 50–1000 bp in length), then extend the new DNA strands. This cycle is repeated to ideally double the number of fragments present in the mixture with each thermocycle. The final amount of amplified DNA fragment produced is dependent on the number of cycles, the number of starting copies of DNA template, and the efficiency of the reaction. Each step of the PCR is temperature-dependent, thus, accurate control of the solution temperature is very important. In the denaturing step, the temperature is raised to approx 95°C where the double-stranded template DNA denatures into single strands. If a high enough temperature is not reached, the DNA will not denature, but too high of a temperature in this step can inactivate the polymerase enzyme; even though it is more heat stable than normal DNA polymerases, Taq polymerase will still degrade in less than 1 h if held at 94°C. Typically, the first denaturing step is maintained for a longer time than subsequent cycles to ensure complete melting of any supercoiled DNA. The temperature is then lowered to within the range of 48 to 74°C depending on the sequence and length of the primers selected; this allows the single-stranded primers to anneal to the single strands of DNA at the appropriate locations. The annealing temperature of a particular target must be optimized empirically using the melting temperature of the primers as a starting point. The optimal annealing temperature is a compromise between the high specificity achieved at higher temperatures and the high percentage of annealing achieved at lower temperatures. If the optimal annealing temperature is not achieved, the specificity of amplification is compromised, where either undesired fragments will be amplified (annealing temperature too low) or not enough products will be formed (annealing temperature too high). After primer annealing, the oligonucleotides are extended through addition of deoxynucleotides by the DNA polymerase using the target DNA sequence as the template. This step is normally performed at the optimal enzymatic temperature (~72°C), but the enzyme is active over a range of temperatures. The dwell time at this step can be short because of the speed of the polymerase enzyme, but it can be extended if long amplicons are being amplified. To achieve sufficient amplification, the whole three-step

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Fig.1. A single thermal cycle for a conventional polymerase chain reaction (PCR) carried out in a heating block. The block temperature (dotted trace) is compared with the actual solution temperature (solid trace). Note that the solution temperature lags behind the block because of heat transfer through the polypropylene tubes. (Reprinted in part with permission from ref. 13. © 2000 American Chemical Society.)

cycle is normally repeated for 20–45 cycles. Often the extension time of the final cycle is held longer to ensure full extension of any incomplete fragments. 1.1. Conventional PCR PCR is typically performed in a commercially available thermocycler that consists of a heating block with up to 96 wells. The PCR mixture is placed in thin wall polypropylene tubes that fit securely into the wells in the block; more recent instruments are also designed for use of microtiter plates in place of the tubes. In the traditional cyclers, the denaturing and annealing steps use dwell times in the range of 5 to 180 s, whereas the extension dwell times are normally tens of seconds to minutes. These times are not limited by the biochemistry of the reaction, but rather by the physics of the process. The required temperature of the mixture for each step is attained by changing the temperature of the heating block in the instrument. Limited by the volume of the solution and heat transfer through the tube wall, the temperature of the solution takes longer to reach the same temperature as the block. Figure 1 shows a typical temperature cycle experienced by the block of a conventional PCR thermocycler where a lag in the reaction solution temperature is observed. The speed of the reaction is, therefore, limited both by the time it takes to change the temperature of the block and the dwell time required for the solution to achieve the proper temperature.

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Decreased thermocycling times have been reported in traditional thermocyclers (2). Though actual solution temperatures might not have been achieved in this system, the authors showed successful amplification for the particular DNA targets chosen. Denaturation of DNA was shown to be completed at temperatures as low as 88°C (2), thus long dwell times at 94°C were not necessary. In the same way, driven by the large excess of primer, DNA annealing was reduced to less than 1 s; as a further benefit, short annealing times have been shown to reduce the amount of mispriming, which leads to nonspecific product (3). The extension step is limited by the enzymatic rate of the polymerase, but at 35–100 nucleotides/s (3), only a few seconds of reaction time are needed near the extension temperature. In addition to time concerns, the cost of the PCR is also an issue, particularly when high-throughput processing is desired. Reaction volumes in polypropylene tubes of traditional thermocyclers are normally in the 25 to 50 µL range; this uses significant amounts of the primers and the Taq polymerase enzyme to achieve the necessary concentrations. Smaller volume (~5 µL) PCR is now possible in traditional cyclers, but significant cost savings could be achieved by reducing the volumes even further. 1.2. Microchip PCR Wilding et al. (4) first demonstrated translation of PCR to a chamber in a microchip device, using silicon/glass hybrid devices that held 5–10 µL of reaction mixture. Though the silicon provided rapid heat transfer, each cycle of PCR was approx 3 min long. The additional thermal mass of the copper block heater and Peltier stage was largely responsible for these slower cycle times. One other issue that arose in the development of microchip PCR was the necessity to coat the chamber walls to prevent adsorption of either the enzyme or the DNA to the silicon or silica surface. This initial work was followed by a subsequent study that investigated the effect of surface passivation on microchip amplification (5). It was found that silanization of the microchamber surface using a covalent coating method provided the best results. A number of other microchip PCR devices and designs have been investigated since this initial work, most notable of which is the flow-through method reported by Kopp et al. in Science (6). All of the currently available microchip PCR methods are detailed in a recent review by Krika and Wilding (7). These authors also report on the types of surface coatings, both dynamic and static, that have been utilized in microchip PCR. One of the benefits of the microchip amplification procedure is the ability to integrate additional processing or analysis steps directly into the same device. Waters et al. (8) performed cell lysis, multiplex PCR amplification, and electrophoretic sizing on a single microchip device. The entire microchip was placed

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into a commercial thermocycler with individual steps of 94°C for 2 min, 37°C for 3 min, and 72°C for 4 min (24 total cycles). The benefits of rapid microscale PCR were unrealized, for the temperature transition times were limited by the thermal mass of the heating block. Burns et al. (9) have approached this problem from an engineering perspective, creating a sophisticated device for amplification and analysis. A resistive heating region with temperature sensors, a sample loading region, and a gel-based separation region were fabricated into a single microdevice; a 106-bp fragment of DNA was successfully amplified and separated. Lagally et al. (10) have developed a device with microfabricated resistance heaters, resistance temperature sensors, and PCR chambers seamlessly connected to electrophoretic separation channels. Valves and hydrophobic vents are used to prevent evaporation of the PCR solution and for fluidic control. Successful sex determination using a multiplex PCR reaction from human genomic DNA was demonstrated in less than 15 min. These two devices are complex and costly to fabricate, however, making them less amenable for large-scale manufacturing or disposability. Development of possible disposable microdevices that contain integrated functionality has been reported by Koh et al. (11) who designed poly (cyclic olefin) devices that incorporated valves and a separation channel, but manufacturing of these devices was still a complex endeavor. 1.3. Infrared Heating One method for reducing the cost of PCR microdevices is to eliminate the need for on-chip heating components. Remote heating of small volume PCR solutions in rectangular glass capillaries using an inexpensive tungsten lamp was first shown by Oda et al. (12). The method relied on direct heating of the solution by excitation of the vibrational bands of the water by adsorption of infrared (IR) radiation from the lamp. This drastically reduced the heat transfer problem associated with heating because the reaction solution was selectively heated. Cooling was performed by flowing a stream of nitrogen past the microcontainers to enhance the rate of heat removal from the container, and thus from the solution. This methodology reduced the total time for one cycle to 17 s, with individual dwell times of 2 s at 94°C, 2 s at 54°C, and 4 s at 72°C. Further efforts showed that decreasing the volume of the reaction to 160 nL increased the efficiency of the PCR (13) and also allowed faster cycling times (Fig. 2). Using a two-temperature protocol recommended by the manufacturer for specific amplification of a 500 bp fragment from a λ-phage DNA template, as few as 10 cycles produced an adequate mass of DNA in less than 12 min for analysis using capillary electrophoresis (CE) with laser-induced fluorescence (LIF) detection. Giordano, et al. (14) applied the IR-mediated heating technique to microchips, performing PCR amplifications in chambers in polyimide microdevices. Because

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Fig. 2. In only 10 thermal cycles, PCR amplification of a 500-bp fragment of λ-DNA was achieved in less than 12 min with only 593 starting copies. This amplification, when compared with conventional thermocycling, illustrates the high efficiency possible with the use of nanoliter scale volumes and exclusive heating of the solution. (Reprinted in part with permission from ref. 13. © 2000 American Chemical Society.)

polyimide is transparent in the 600- to 3000-nm range, the IR radiation again selectively heated the solution, and the low thermal mass of the chips provided rapid cooling even without forced convection. Heating and cooling rates as high as 10°C/s and short dwell times resulted in cycles that required only 12 s to complete. Adequate amounts of PCR product were observed using LIF detection after 15 cycles, a process taking only 240 s. Surface passivation was also required with these devices, with polyethylene glycol included in the PCR mixture to dynamically coat the chip surface. Polyimide does not make a good substrate for integration of additional processing steps owing to incompatibilities with existing separation and detection techniques however, so the IR method was transferred to glass microdevices. With the much greater thermal mass of the glass chips and the ability of glass to dissipate heat, the heating and cooling rates were reduced, but amplification could still be performed in approx 20 min. Figure 3 shows a thermocycling profile for a glass microdevice using IR-mediated heating along with a microchip electrophoretic analysis of PCR product for amplification of the 275bp invasive A (invA) gene from Salmonella typhimurium (primers designed inhouse). In this example, separation and detection were performed on separate microdevices, but PCR product could also be directly analyzed in an integrated

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Fig. 3. PCR amplification of the target 275-bp invasive A (invA) gene of Salmonella typhimurium DNA using primers designed in-house. (A) Three-temperature thermocycling was carried out as follows: 35 cycles at 95°C for 3 s, 64°C for 8 s, and 75°C for 3 s, with a final extension at 75°C for 60 s (total time ~21 min). (B) Separation was carried out by microchip gel electrophoresis with sizing markers of 15 and 1500 bp to confirm the presence of the 275-bp amplicon.

electrophoretic separation channel on the same device (15). Surface passivation in these devices relied on an absorbed epoxy poly(dimethylacrylamide) (EPDMA) coating that was validated for glass devices (16) with bovine serum albumin (BSA) added to the PCR mixture for additional passivation. This chapter details the IR-mediated amplification method utilized in these publications, but focuses on simple glass devices containing only the PCR chamber and a thermocouple reference chamber.

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2. Materials 2.1. Fabrication of Microdevices 1. Borofloat glass plates 1.1-mm thick, coated with chrome and photoresist (Nanofilms, Westlake, CA). 2. Borofloat glass for cover plates 1.1-mm thick (S.I. Howard Glass, Worcester, MA) (see Note 1). 3. Flood ultraviolet (UV) light source (OAI Associates, Milipitas, CA). 4. Mask with image of device (Fig. 4) (see Note 2). 5. Photoresist developer AZ 400K (Clariant Corp, Sunnyvale, CA). 6. Chromium etchant CR-7S (Cyantek Corp, Fremont, CA). 7. Photoresist stripper AZ 300T (Clariant Corp). 8. Hydrofluoric acid (HF). 9. Nitric acid (HNO3). 10. Diamond-tipped drill bits (Crystalite Corp, Lewis Center, OH). 11. Ceramic plates. 12. Colloidal graphite (Renite S-24, Columbus, OH).

2.2. Microchip Reservoirs 1. Sylgard 184 silicone elastomer base and curing agent (Dow Corning Corp, Midland, MI), both stored below 32°C. 2. Vacuum desiccation chamber.

2.3. Coating Polymer Preparation 1. 2. 3. 4. 5.

N,N-dimethylacrylamide (Aldrich). Allylglycidyl ether (Aldrich). TEMED (Aldrich). Ammonium persulfate (APS) solution (40% w/v) in distilled water. Dialysis tubing.

2.4. Preparation of the Microchip 1. 1 M NaOH. 2. Autoclaved Nanopure water (Barnstead International, Dubuque, IA). 3. 2 mg/mL BSA (Aldrich).

2.5. Temperature Detection 1. Miniature type-T copper-constantan thermocouple (T240C, Physitemp Instruments, Clifton, NJ). 2. Thermocouple amplifier (TAC-386-TC; Omega Engineering, Inc., Stamford, CT) with output of 1 mV/°C.

2.6. IR-Mediated Thermocycler 1. 5-V/12-V DC power supply (Power-One, Camarillo, CA). 2. 8-V, 50-W Tungsten filament lamp (General Electric, Cleveland, OH). 3. Solenoid valve with a room-temperature compressed air source (such as a nitrogen tank) and appropriate tubing.

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Fig. 4. Positive mask design for a PCR microchip including (A) a reference chamber for the thermocouple and (B) a PCR chamber for DNA amplification. Void regions were also etched in proximity to the chambers to reduce thermal losses to the glass. The mask negative should be used for ultraviolet exposure. 4. Circuitry for active control of lamp and solenoid (see Note 3). 5. LabVIEW application to control the switching circuitry (see Note 4). 6. Gold-mirrored surface to enhance heating (Edmund Industrial Optics, Barrington, NJ).

2.7. Preparation of the PCR Mixture 1. 2. 3. 4.

25 mM MgCl2, stored at –20°C. 10X PCR buffer: 100 mM Tris, 500 mM KCl, stored at –20°C (see Note 5). 100 mM dNTPs: dATP, dGTP, dCTP, dTTP, store at –20°C. 20 µM of forward and reverse primers complimentary to the end regions of the DNA fragment being amplified. 5. 5000 U/mL Taq polymerase. 6. 2 mg/mL BSA. 7. Template DNA containing the fragment to be amplified. The specific DNA fragment will dictate the amount of starting material that will be required.

2.8. Loading the Microchip and Thermocycling 1. Molecular biology grade mineral oil.

3. Methods 3.1. Fabrication of Microdevice 1. The borofloat glass with chrome and photoresist are exposed to the UV source through the mask negative for 5 s (see Note 6). 2. The exposed photoresist is removed using developer then the resist hard-baked at 110°C for 30 min. 3. The exposed chrome is removed using chromium etchant.

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Fig. 5. Image of a completed microdevice for PCR amplification. The channels were filled with blue dye for visualization. 4. The glass is etched using a solution of HF:HNO3 :H2O (50:14:36), etching at a rate of 3 µm/min. 5. The glass is etched to 150-µm deep. 6. The remaining photoresist is removed using stripper. 7. The chrome is removed using chromium etchant. 8. Top plates are fabricated by drilling reservoir holes that will line up with the ends of the PCR chamber using a diamond-tipped drill bit (1.1-mm diameter). 9. Bottom and top plates are cut to size then cleaned with an ammonia-based window cleaner (Windex® window cleaner). 10. The glass plates are pressed together, placed between graphite-coated ceramic plates, and placed in a high-temperature furnace for bonding. 11. The furnace temperature is ramped to 550°C at 8°C/min, then at 3°C/min to 670°C where it is held for 3.5 h before naturally cooling to room temperature to avoid cracking. Figure 5 shows a completed device (see Note 7).

3.2. Microchip Reservoirs 1. Combine silicone elastomer base and curing agent in a 15:1 ratio, mixing well (see Note 8).

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2. Degas the mixure in a vacuum desiccator for 15 min or until no air bubbles remain. 3. Pour degassed mixture into a Petri dish to a depth of approx 5 mm. 4. Bake at 60°C for 2 h, making sure that the dish is level. 5. With a razor blade, cut a piece of the cured poly(dimethylsiloxane) (PDMS) to match the shape of the microchip, and produce the appropriate reservoirs with a hole punch (see Note 9). 6. Thoroughly clean the PDMS and glass microchip surfaces with soap and water, and dry them well with a stream of nitrogen to ensure a tight seal to the glass. 7. Align the PDMS reservoir with the etched reservoirs of the microchip, bring them into contact, and apply light pressure until no air bubbles exist between the two layers. Although unnecessary, heating the chip at this point (~60°C for ~10 min) has been found to enhance the seal. This reservoir should be capable of containing any aqueous solution, and the seal is readily reversible. 8. If the seal is ineffective, simply peel away and repeat steps 6 and 7.

3.3. Preparation of Coating Polymer (17) 1. A 10 mL solution of 0.4 M N,N-dimethylacrylamide, 0.008 M allylglycidyl ether in water is degassed for 10 min. 2. The reaction is initiated using 10 µL of TEMED and 10 µL of APS solution added below the surface, and allowed to react for 24 h at room temperature. 3. The reaction mixture is dialyzed for 4 h against three changes of distilled water to remove any unreacted monomer. 4. The mixture is lyophylized to obtain a solid, then resuspended in water at a concentration of 0.2% (w/v).

3.4. Preparation of the Microchip 1. 2. 3. 4. 5.

Rinse the chamber with 1.0 M NaOH for 10 min to prepare for passivation. Flush for 10–20 min with 0.2% EPDMA to passivate the chamber (see Note 10). Rinse with distilled, autoclaved water for 20 min. Rinse with BSA for 5 min to enhance the EPDMA passivation. Rinse with distilled, autoclaved water for 5 min.

3.5. Temperature Detection 1. If needed, carefully sand the end of the miniature thermocouple with fine sandpaper until it will fit into the reference chamber. 2. Gently thread the thermocouple into the central region of the reference chamber where the lamp will be focused (see Note 11). 3. Secure the thermocouple in place by taping to the chip.

3.6. IR-Mediated Thermocycler 1. Build the appropriate electrical circuit box to allow computer control of the heating lamp and solenoid air valve (see Note 12).

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2. Build a chip holder that positions the microchip reaction chamber in the focus of the lamp and in the stream of the compressed air source (see Note 13).

3.7. Preparation of the PCR Mixture (25 µL total volume) (see Note 14) 1. Combine the following reagents to make the “PCR mixture” (see Note 15): 16.6 µL distilled, autoclaved water. 3 µL 25 mM MgCl2 (see Note 5). 2.5 µL 10X PCR buffer. 0.05 µL 100 mM dATP. 0.05 µL 100 mM dTTP. 0.05 µL 100 mM dGTP. 0.05 µL 100 mM dCTP. 0.2 µL 20 µM forward primer. 0.2 µL 20 µM reverse primer. 0.3 µL 2 mg/mL BSA. 2 µL DNA at the appropriate concentration for the application (water is used as a negative control) (see Note 16). 2. To each 25-µL aliquot of the previously mentioned mixture, add 0.6 µL of Taq polymerase. a. b. c. d. e. f. g. h. i. j. k.

3.8. Loading the Microchip and Thermocycling 1. Making sure the passivated PCR chamber is empty, use a pipet to add about 20 µL of the PCR mixture to one of the reservoirs and allow capillary action to pull it through the chamber and into the opposing reservoir (see Note 17). 2. Fill the reference chamber in the same manner with PCR buffer. 3. Cover the solution in all reservoirs with just enough mineral oil to prevent evaporation. 4. Place the microchip into the holder above the focal point of the IR heating source in a position to promote equal heating of the entire PCR chamber and the reference chamber 5. Place the mirrored surface above the heated region to increase the heating rate. 6. The denaturing/annealing/extension temperatures appropriate for the gene to be amplified are entered in the program (see Notes 18 and 19). Figure 3 shows an example of thermocycling along with the conditions used.

4. Notes 1. Borofloat glass for etching and cover plates can also be purchased in 0.7-mm thickness, which can be used to fabricate these devices. The thinner glass allows an increased speed of thermocycling, but results in more fragile microchips. 2. Photographic negative masks can be prepared from AutoCAD files of the microdevice design by commercial printers at approx 6 dpi resolution using lithographic printing. Metal masks can also be designed for this purpose. 3. Only beginner-level knowledge of electronics is necessary for this purpose. The lamp and solenoid should be connected to solid-state relays, which can feed the

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5.

6. 7.

8.

9.

10.

11.

12. 13.

14.

15.

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appropriate voltages from the power supply. The relays should be connected to a PC outfitted with an analog-to-digital (A/D) converter board to allow computer control. A proportional integral differential algorithm is best suited for thermal control of PCR. This type of algorithm will help the system avoid overshooting the temperatures needed for PCR. The LabVIEW application should use the input from the thermocouple to actively control the solenoid and solid-state relays, and maintain the temperature of the system at the input values. Note that the 10X PCR buffer can be purchased both with and without MgCl2. If the correct concentration of MgCl2 is included with the 10X PCR buffer, then adding MgCl2 is not necessary. Figure 4 shows a positive image of an example mask pattern for microchip PCR. The negative image should be used for UV exposure. There are multiple chip geometries possible, and many are likely suitable for use with the protocol. This particular straight-channel geometry was chosen simply because the authors have been successful with its use. A 15:1 ratio of elastomer:curing agent has been found to enhance the seal between the PDMS reservoir and the glass microchip when compared with the 10:1 ratio recommended by the manufacturer. To ensure accurate reservoir fabrication, use a printout of the microchip design as a guide. Also, be sure to create a larger reservoir at the reference chamber to ease the installation of the thermocouple. It is important not to rinse with water before the EPDMA passivation. The 1 M NaOH rinse before the coating is used to prepare the surface for polymer adsorption, and rinsing with water will adversely alter the surface. This technique takes time to master, and use of a microscope is helpful. Tweezers can be used to thread the thermocouple into the chamber, but they may cause abrasions to the coating and foul the sensor. Pipet tips were found to be useful for this purpose, with sensor fouling much less likely. Also, miniature thermocouples are very delicate. Caution must be taken when transporting the microchip to different stations if the thermocouple is installed. The fast response time of solid-state relays make them ideal for this application, but any sort of switching circuitry may be used. The chip holder should be constructed from an insulating material to avoid fast heat dissipation. If the holder is too good of a heat sink, the solution in the chamber may not be able to reach the high temperatures or may take too long to reach them. Less than 1 µL is needed to fill the PCR chamber, but the reservoirs contain more to help prevent evaporation. The volume of the solution can be reduced by decreasing the size of the reservoirs to save expensive reagents while still achieving amplification. Allow all PCR reagents except the Taq polymerase to thaw to room temperature. Because Taq is thermally sensitive, it should be kept at –20°C and should be the last reagent added to the reaction. The thermocycling should be initiated within 5 min of enzyme addition.

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16. Because the surface area-to-volume ratio is drastically increased in microdevices, some DNA and enzyme may still adsorb to the glass surface, even with the passivation. Two to five nanograms of genomic DNA is a typical amount included in a single reaction mixture (25 µL). 17. Be sure to allow room in the reservoirs for the mineral oil when filling with the PCR mixture. Overflowing the reservoirs with mineral oil can provide a leakage path while thermocycling. If the mineral oil leaks away, the reaction mixture will evaporate. 18. The annealing temperature is dependent on the primer sequence; primers chosen for PCR are dependent on the DNA fragment being amplified. 19. Because the overall reaction environment is different in glass microchips than in traditional thermocyclers, some adjustments to the typical temperature protocols may be required for optimal amplification.

References 1. Saiki, R. K., Scharf, S., Faloona, F., et al. (1985) Enzymatic amplification of betaglobin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230, 1350–1354. 2. Mai, M., Grabs, R., Barnes, R. D., Vafiadis, P., and Polychronakos, C. (1998) Shortened PCR cycles in a conventional thermal cycler. Biotechniques 25, 208–210. 3. Wittwer, C. T. and Garling, D. J. (1991) Rapid cycle DNA amplification: time and temperature optimization. Biotechniques 10, 76–83. 4. Wilding, P., Shoffner, M. A., and Kricka, L. J. (1994) PCR in a silicon microstructure. Clin. Chem. 40, 1815–1818. 5. Wilding, P., Shoffner, M. A., Cheng, J., Huichia, G., and Kricka, L. J. (1995) Thermal cycling and surface passivation of micromachined devices for PCR. Clin. Chem. 41, 1367, 1368. 6. Kopp, M. U., Mello, A. J., and Manz, A. (1998) Chemical amplification: continuousflow PCR on a chip. Science 280, 1046–1048. 7. Kricka, L. J. and Wilding, P. (2003) Microchip PCR. Anal. Bioanal. Chem. 377, 820–825. 8. Waters, L. C., Jacobson, S. C., Kroutchinina, N., Khandurina, J., Foote, R. S., and Ramsey, J. M. (1998) Microchip device for cell lysis, multiplex PCR amplification, and electrophoretic sizing. Anal. Chem. 70, 158–162. 9. Burns, M. A., Mastrangelo, C. H., Sammarco, T. S., et al. (1996) Microfabricated structures for integrated DNA analysis. Proc. Natl. Acad. Sci. USA 93, 5556–5561. 10. Lagally, E. T., Emrich, C. A., and Mathies, R. A. (2001) Fully integrated PCRcapillary electrophoresis microsystem for DNA analysis. Lab on a Chip 1, 102–107. 11. Koh, C. G., Tan, W., Zhao, M. Q., Ricco, A. J., and Fan, Z. H. (2003) Integrating polymerase chain reaction, valving, and electrophoresis in a plastic device for bacterial detection. Anal. Chem. 75, 4591–4598.

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12. Oda, R. P., Strausbauch, M. A., Huhmer, A. F., et al. (1998) Infrared-mediated thermocycling for ultrafast polymerase chain reaction amplification of DNA. Anal. Chem. 70, 4361–4368. 13. Huhmer, A. F. and Landers, J. P. (2000) Noncontact infrared-mediated thermocycling for effective polymerase chain reaction amplification of DNA in nanoliter volumes. Anal. Chem. 72, 5507–5512. 14. Giordano, B. C., Ferrance, J., Swedberg, S., Huhmer, A. F., and Landers, J. P. (2001) Polymerase chain reaction in polymeric microchips: DNA amplification in less than 240 seconds. Anal. Biochem. 291, 124–132. 15. Ferrance, J. P., Wu, Q., Giordano, B., et al. (2003) Developments toward a complete micro-total analysis system for Duchenne muscular dystrophy diagnosis. Analytica Chimica Acta 500, 223–236. 16. Giordano, B. C., Copeland, E. R., and Landers, J. P. (2001) Towards dynamic coating of glass microchip chambers for amplifying DNA via the polymerase chain reaction. Electrophoresis 22, 334–340. 17. Chiari, M., Cretich, M., Damin, F., Ceriotti, L., and Consonni, R. (2000) New adsorbed coatings for capillary electrophoresis. Electrophoresis 21, 909–916.

Index

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Index A Amperometric detection, capillary electrophoresis, 92, 93 externally positioned off-chip electrode alignment, advantages, 98, 99 applications, 103, 105 micromanipulated electrode, 99, 101, 107 screen-printed approach, 99 injection, 88, 89, 91, 92, 105, 106 integrated on-chip electrode alignment, applications, 101–103 decoupled detection, 97, 98 end-channel detection, 95, 107 in-channel detection, 95–97 working electrode potential and alignment, 94, 107 materials, 87, 88 popularity, 86 principles, 86, 87 voltage separation, 93, 94

analytes and applications, 120–124 buffer systems, 118, 120 electrode manufacturing, 114–118, 124 electronics, 118 materials, 114 Current monitoring, see Electroosmotic flow D Destaining, microchips for protein separation, 146–148 Direct-write, see Laser ablation DNA amplification, see Polymerase chain reaction DNA separation, historical perspective, 129, 130 lab-on-a-chip device separation, materials, 131, 133–135, 140–142 planar microfluidic chips, 135–138 principles, 130, 131 sipper microfluidic chips, 138–142

B, C Biotinylation, surface modification, 54, 55 Bonding, see Glass bonding Capillary electrochromatography (CEC), principles, 2–4 Capillary zone electrophoresis, peptide analysis, 165, 166, 179, 180 CEC, see Capillary electrochromatography Cell analysis, see Single cell analysis Conductivity detection, advantages and limitations, 113, 114

E Electrochemical detection, see also Amperometric detection; Labon-a-chip, modes, 86 peptide detection, 176–178, 180 Electrochromatography, peptide analysis, 169–171 Electroosmotic flow (EOF), measurement, current monitoring method, materials, 192 overview, 190 technique, 194–199

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234 flow imaging, 190, 191 neutral marker method, materials, 191, 192 overview, 189, 190 technique, 193, 194, 196–198 polyelectrolyte multilayers, 61 principles, 2–4, 187–189 reproducibility, 188 Electrospray coupling, see Mass spectrometry Enzyme assay, peptide analysis, 172–174 EOF, see Electroosmotic flow F, G Faraday’s law, 87 Fast prototyping, see Hot embossing Glass bonding, DNA separation microchips, 23–25, 34 electrospray-coupled microchips, 72, 73, 78 H Hot embossing, materials, 38, 45 microchannel fabrication, 43, 44 poly(dimethylsiloxane) microchip, 33, 34 polymer embossing, 40–42 principles, 37, 38 silicon template micromatching, 39, 40 I, J Immunoassay, peptide analysis, 171, 172 Infrared heating, microchip polymerase chain reaction, 221–223, 227, 229 Instantaneous current, equation, 87 Isoelectric focusing, peptide analysis, 167–169 Isotachophoresis, peptide analysis, 167 Jurkat cell, see Single cell analysis L Lab-on-a-chip (LOC), device design, 14

Index DNA separation, materials, 131, 133–135, 140–142 planar microfluidic chips, 135–138 principles, 130, 131 sipper microfluidic chips, 138–142 microfabrication, bottom substrate processing, 17–20, 34 electrode integration, 25, 26 glass-to-glass bonding, 23–25, 34 photolithography, 16, 17 photomask development, 14, 16, 34 top substrate processing, 20–23 protein separation, advantages, 157, 158 denaturation of samples, 150, 151 destaining, 146–148 materials, 148–150, 154 planar microfluidic chips, 151, 152, 154, 155, 157 sipper microfluidic chips, 152–155, 157 Laser ablation, materials, 38, 45 poly(dimethylsiloxane) microchip, 34 principles, 38 technique, 42–44 Laser-induced fluorescence (LIF), overview of detection, 85, 86 single cell analysis, data collection and analysis, 213 setup, 209 LIF, see Laser-induced fluorescence LOC, see Lab-on-a-chip M Mass spectrometry (MS), advantages in capillary electrophoresis detection, 68 electrospray coupling of microchip capillary electrophoresis, chip fabrication, access hole drilling, 73, 74, 78 fittings, 74

Index glass bonding, 72, 73, 78 photolithography, 71, 72, 78 wet chemical etching, 72, 78 coating of microchips, 74, 79 interfacing and operation, 75–79 materials, 70, 71 overview, 68–70 spray tip preparation, 75 peptide detection, 178, 179 Mass, calculation for analyte, 3 MEKC, see Micellar electrokinetic chromatography Micellar electrokinetic chromatography (MEKC), peptide analysis, 166, 167 principles, 2, 3 Microchip capillary electrophoresis, applications, DNA separation, 7 overview, 2 protein analysis, 7 small molecule analysis, 8 detection, 6, 7 historical perspective, 1, 2 injection, 5 microchip construction, 4, 5 separation, 6 theory, 2–4 Microdialysis, peptide samples, 163 MS, see Mass spectrometry N–P Net charge, calculation for analyte, 3 Neutral marker, see Electroosmotic flow PCR, see Polymerase chain reaction PDMS microchip, see Poly(dimethylsiloxane) microchip Peptide analysis, applications, 159, 160 considerations, 160 microchip electrophoresis, capillary zone electrophoresis, 165, 166, 179, 180 detection,

235 electrochemical detection, 176–178, 180 fluorescence, 175, 176 mass spectrometry, 178, 179 electrochromatography, 169–171 enzyme assay, 172–174 immunoassays, 171, 172 injection, 164 isoelectric focusing, 167–169 isotachophoresis, 167 micellar electrokinetic chromatography, 166, 167 multi-dimensional separations, 174 principles, 160–162 sample preparation, digestion of proteins, 164, 179 microdialysis, 163 preconcentration and desalting, 162, 163 substrate selection, 161, 162, 179 Photolithography, electrospray-coupling chip fabrication, 71, 72, 78 lab-on-a-chip, 16, 17 poly(dimethylsiloxane) microchip, alignment and exposure, 30 development, 31, 32, 34, 35 hard bake, 32, 35 mask preparation, 28, 29 photoresist deposition, 29, 34 postexposure baking, 31 soft bake, 30 water surface preparation, 29, 34 Planar microfluidic chip, DNA separation, 135–138 protein separation, 151, 152, 154, 155, 157 Poly(dimethylsiloxane) (PDMS) microchip, advantages, 27, 57 coating of glass microchips, 51, 54, 55 fabrication, assembly, irreversible sealing, 33, 59 reversible sealing, 33, 59

236 checklist, 33 direct-write, 34 fast prototyping, 33, 34 materials, 28 overview, 27, 28 photolithography, alignment and exposure, 30 development, 31, 32, 34, 35 hard bake, 32, 35 mask preparation, 28, 29 photoresist deposition, 29, 34 postexposure baking, 31 soft bake, 30 water surface preparation, 29, 34 substrate characteristics and preparation, 32 polyelectrolyte multilayer, see Polyelectrolyte multilayer Polyelectrolyte multilayer (PEM), electroosmotic flow measurements, 61 materials, 58, 59 noncovalent coating, 59, 60 polyelectrolyte types, 63 rationale, 57, 58 stability and reproducibility of coating, 61, 62 surface modification, 50, 53, 55 Polymerase chain reaction (PCR), conventional thermocycling, 219, 220 historical perspective, 217, 218 microchip polymerase chain reaction, coating polymer preparation, 227 infrared heating, 221–223, 227, 229 loading and thermocycling, 228, 230 materials, 224, 225, 228, 229 microchip fabrication, 225, 226, 227, 229 microchip reservoirs, 226, 227, 229

Index overview, 220, 221 reaction mixture preparation, 228, 229 temperature detection, 227, 229 principles, 218, 219 Poly(methylmethacrylate) devices, amine termination, 51, 54, 55 Protein separation, gel electrophoresis overview, 145, 146 lab-on-a-chip device separation, advantages, 157, 158 denaturation of samples, 150, 151 destaining, 146–148 materials, 148–150, 154 planar microfluidic chips, 151, 152, 154, 155, 157 sipper microfluidic chips, 152–155, 157 R, S Reservoir, microchip reservoirs for polymerase chain reaction, 226, 227, 229 Sealing, poly(dimethylsiloxane) microchip assembly, irreversible sealing, 33, 59 reversible sealing, 33, 59 thermal bonding, 43, 44, 45 Single cell analysis, microfluidic devices, advantages, 204, 205 channel wall coating and durability, 210, 211, 213, 214 imaging of chip and cell, 212, 213 injection efficiency measurement, 213 Jurkat cell culture, 211 laser-induced fluorescence detection, data collection and analysis, 213 setup, 209 loading of cells, 211

Index materials, 205, 206, 213, 214 microchip, bonding, 208, 209, 214 design and operation, 206, 207 glass etching, 208, 214 transport and lysis of cells, 211, 212, 214 overview of techniques, 203, 204 Sipper chip, DNA separation, 138–142 protein separation, 152–155, 157 Surface modification, amine termination of poly(methylmethacrylate) devices, 51, 54, 55 biotinylation, 54, 55 materials, 51–53

237 poly(dimethylsiloxane) coating of glass microchips, 51, 54, 55 polyelectrolyte multilayer, 50, 53, 55 rationale, functionalization of surfaces, 51 practical effects, 49, 50 surface property improvement, 50, 51 ultraviolet graft polymerization, 53 T–V Temperature detection, microchip polymerase chain reaction, 227, 229 Ultraviolet graft polymerization, surface modification, 53 Velocity, formula, 3