Integrated Microfabricated Biodevices

Cover artwork by courtesy of Nanogen, Inc., San Diego, California, with permission. ISBN: 0-8247-0606-4 This book is printed on acid-free paper. Headquarters Marcel Dekker, Inc. 270 Madison Avenue, New York, NY 10016 tel: 212-696-9000; fax: 212-685-4540 Eastern Hemisphere Distribution Marcel Dekker AG Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland tel: 41-61-261-8482; fax: 41-61-261-8896 World Wide Web http:/ /www.dekker.com The publisher offers discounts on this book when ordered in bulk quantities. For more information, write to Special Sales/Professional Marketing at the headquarters address above. Copyright  2002 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Current printing (last digit): 10 9 8 7 6 5 4 3 2 1 PRINTED IN THE UNITED STATES OF AMERICA

Preface

Interdisciplinary science and technologies have converged in the last few years to create exciting challenges and opportunities that involve a new generation of integrated microfabricated devices. These new devices are referred to as DNA chips, Lab-on-a-Chip, microcapillary electrophoresis systems, microanalytical systems, and biosensors. Their development involves both established and evolving technologies, including microlithography, micromachining, microelectromechanical systems (MEMS technology), microfluidics, and nanotechnology. The development of these new devices and systems requires a high level of interaction and cooperation among engineers, computer scientists, materials scientists, chemists, molecular biologists, geneticists, and clinical scientists. Applications for this novel ‘‘synergized’’ technology will include genetic analysis, clinical chemistry (particularly DNA and immunodiagnostics), drug discovery, combinatorial chemistries, industrial process control, and portable, hand-held analytical instrumentation. The first aim of this book is to provide a good overview of the key devices (DNA chips, Lab-on-a-Chip, etc.) and the basic interdisciplinary technologies (microfabrication, MEMS, microfluidics, etc.). The second aim is to give the reader a better understanding of how to utilize these interdisciplinary technologies and determine which will provide appropriate technical solutions to problems perceived as being limited to their own discipline. Our contributors have identified important aspects of these broad interdisciplinary technologies that are particularly relevant to successful development of novel devices or systems. For example: DNA chips and arrays may be highly dependent on certain microfabrication or micromachining techniques and processes, whereas Lab-on-a-Chip and microcapillary electrophoresis systems are more dependent on so-called microfluidic processes. Successful development of a given device or system requires overcoming a variety iii

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of problems. While this sounds self-evident, and generally does not represent a problem within a given discipline, it is indeed a problem in highly interdisciplinary areas. Thus, readers need to obtain a firm grasp of the underlying base technologies and/or applications to make better decisions about their own areas of research and development. The information presented here will help the reader make decisions as to whether a particular problem (often perceived as fundamental within a discipline) can be solved by using another technology. Finally, we would also like the reader to understand that as novel as these applications sound, the door is just opening to even more exciting advancements. Because of this, we complete this volume with a look at the so-called area of nanotechnology, which may herald the advent of molecular electronics and nanocomputing systems. Because of the highly interdisciplinary nature of this book, it should be of interest and value to a relatively large audience. The text will serve a wide range of academic and industrial scientists and engineers, including molecular biologists, geneticists, and microbiologists; clinical and diagnostic scientists; nucleic acid, organic, physical, and analytical chemists; electronic and mechanical engineers; computer scientists and programmers; and physicists and mathematicians. This volume will also be valuable to many entrepreneurial and business people who are in the process of trying to better understand and evaluate these new, fast-moving high-tech areas. Michael J. Heller Andra´s Guttman

Contents

Preface Contributors 1. Toward an Integrated Electrophoretic Microdevice for Clinical Diagnostics Braden C. Giordano, Jerome Ferrance, and James P. Landers 2. Creating a Lab-on-a-Chip with Microfluidic Technologies Anne R. Kopf-Sill, Andrea W. Chow, Luc Bousse, and Claudia B. Cohen 3. Short Tandem Repeat Analysis on Microfabricated Electrophoretic Devices Dieter Schmalzing, Aram Adourian, Lance Koutny, Paul Matsudaira, and Daniel Ehrlich 4. Basic Sample Preparation: DNA Separation and Purification Rama Ramanujam, Wendy Sacks, and Jie Kang 5. Multiplexed Integrated Online Sample Preparation for DNA Sequencing and Genetic Typing Edward S. Yeung 6. Interfacing Capillaries to Microseparation Devices for Sample Introduction Steven W. Suljak and Andrew G. Ewing

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Contents Electric-Field-Mediated Separation of DNA Fragments on Planar Microgels Andra´s Guttman Development of Battery-Powered, Portable Instrumentation for Rapid PCR Analysis Phillip Belgrader, Bill Benett, Dean Hadley, Ray Mariella, Jr., M. Allen Northrup, Jim Richards, Paul Stratton, Shanavaz Nasarabadi, and Fred Milanovich Practical Aspects of Sheath Flow Fraction Collection in Capillary Electrophoresis Odilo Mu¨ller and Hirofumi Suzuki Active Microelectronic Array Systems for DNA Hybridization, Genotyping, Pharmacogenomic, and Nanofabrication Applications Michael J. Heller, Eugene Tu, Robert Martinsons, Richard R. Anderson, Christian Gurtner, Anita H. Forster, and Ron Sosnowski The Flow-thru Chip: A Miniature, Three-Dimensional Biochip Platform Adam Steel, Matt Torres, John Hartwell, Yong-Yi Yu, Nan Ting, Glenn Hoke, and Hongjun Yang

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DNA Arrayed on Plastic Devices Robert S. Matson

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Microfluidic Devices Fabricated by Polymer Hot Embossing Holger Becker

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Microfabricated Reactor Technology Tibor Chova´n and Andra´s Guttman

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Approaches to Miniaturized High-Throughput Screening of Chemical Libraries Damien Dunnington, Zhuyin Li, Alan Binnie, and Henning Vollert

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16. Integrated Microsystems in Clinical Chemistry: Constraints and Consequences Michael Bell

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Index

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Contributors

Aram Adourian Whitehead Institute for Biomedical Research, Cambridge, Massachusetts Richard R. Anderson Nanogen, Inc., San Diego, California Holger Becker Mildendo GmbH, Jena, Germany Phillip Belgrader Microfluidic Systems Inc., Berkeley, California Michael Bell Beckman Coulter, Inc., Brea, California Bill Benett Lawrence Livermore National Laboratory, Livermore, California Alan Binnie Aventis, Bridgewater, New Jersey Luc Bousse Caliper Technologies Corp., Mountain View, California Claudia B. Cohen Caliper Technologies Corp., Mountain View, California Tibor Chova´n University of Veszpre´m, Veszpre´m, Hungary Andrea W. Chow Caliper Technologies Corp., Mountain View, California Damien Dunnington Aventis, Bridgewater, New Jersey Daniel Ehrlich Whitehead Institute for Biomedical Research, Cambridge, Massachusetts Andrew G. Ewing Pennsylvania State University, University Park, Pennsylvania Jerome Ferrance University of Virginia, Charlottesville, Virginia Anita H. Forster Nanogen, Inc., San Diego, California ix

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Braden C. Giordano University of Virginia, Charlottesville, Virginia Christian Gurtner Nanogen, Inc., San Diego, California Andra´s Guttman Torrey Mesa Research Institute, San Diego, California Dean Hadley Lawrence Livermore National Laboratory, Livermore, California John Hartwell Gene Logic, Incorporated, Gaithersburg, Maryland Michael J. Heller Nanogen, Inc., and University of California, San Diego, San Diego, California Glenn Hoke Gene Logic, Incorporated, Gaithersburg, Maryland Jie Kang QIAGEN GmbH, Hilden, Germany Anne R. Kopf-Sill Caliper Technologies Corp., Mountain View, California Lance Koutny Whitehead Institute for Biomedical Research, Cambridge, Massachusetts James P. Landers University of Virginia, Charlottesville, Virginia Zhuyin Li Aventis, Bridgewater, New Jersey Ray Mariella, Jr. Lawrence Livermore National Laboratory, Livermore, California Robert Martinsons Nanogen, Inc., San Diego, California Robert S. Matson Beckman Coulter, Inc., Fullerton, California Paul Matsudaira Whitehead Institute for Biomedical Research, Cambridge, Massachusetts Fred Milanovich Lawrence Livermore National Laboratory, Livermore, California Odilo Mu¨ller Agilent Technologies Deutschland GmbH, Waldbronn, Germany Shanavaz Nasarabadi Lawrence Livermore National Laboratory, Livermore, California M. Allen Northrup Microfluidic Systems Inc., Berkeley, California Rama Ramanujam QIAGEN GmbH, Hilden, Germany

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Jim Richards Lawrence Livermore National Laboratory, Livermore, California Wendy Sacks QIAGEN GmbH, Hilden, Germany Dieter Schmalzing Whitehead Institute for Biomedical Research, Cambridge, Massachusetts Ron Sosnowski Nanogen, Inc., San Diego, California Adam Steel Gene Logic, Incorporated, Gaithersburg, Maryland Paul Stratton Lawrence Livermore National Laboratory, Livermore, California Steven W. Suljak Pennsylvania State University, University Park, Pennsylvania Hirofumi Suzuki Organon, Ltd., Osaka, Japan Nan Ting Gene Logic, Incorporated, Gaithersburg, Maryland Matt Torres Gene Logic, Incorporated, Gaithersburg, Maryland Eugene Tu Nanogen, Inc., San Diego, California Henning Vollert Aventis, Bridgewater, New Jersey Hongjun Yang Gene Logic, Incorporated, Gaithersburg, Maryland Edward S. Yeung Iowa State University, Ames, Iowa Yong-Yi Yu Gene Logic, Incorporated, Gaithersburg, Maryland

1 Toward an Integrated Electrophoretic Microdevice for Clinical Diagnostics Braden C. Giordano, Jerome Ferrance, and James P. Landers University of Virginia, Charlottesville, Virginia

1.1 INTRODUCTION Recent years have seen much effort invested in developing clinical diagnostic tools that provide both rapid analysis and accurate results. The growing literature on microminiaturized analytical devices has begun to delineate the potential applicability of the microfabricated chip platform for this task. While the enhanced analytical capabilities of microfabricated chip technology for expediting electrophoretic separations have been well established [1–8], the true power of this technology lies in the potential to miniaturize and integrate existing technologies in a manner that allows for sample preparation and analysis to be seamlessly carried out on a single device. Many routine assays carried out in the clinical setting rely on slab gel electrophoresis for detection of DNA and proteins. Capillaries have catalyzed the first step in miniaturizing this process, having been shown to be an effective alternative to slab gel-based methods while reducing analysis times by roughly an order of magnitude [1,9–11]. In addition, the low volumes of reagents consumed by this electrophoretic process 1

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combined with the miniscule sample requirements bolster the advantages it brings to clinical analysis. The disadvantage associated with the capillary format for electrophoretic analysis is that, while slab gel electrophoresis allows for the parallel processing of a number of samples simultaneously, capillary-based separations usually must be executed one by one. Consequently, any gain through reduction of analysis time is lost by serial analysis. While this yields an overall process time for a batch of samples that is roughly the same with both methods, capillary electrophoresis (CE) still has brought a number of benefits including semiautomation and online detection, both of which are advantageous in the clinical laboratory [12]. Moreover, newer CE instrumental platforms that incorporate multiple capillaries for the concurrent processing of a large number of samples [13–17] have solved the serial analysis problem. While CE can reduce analysis times by orders of magnitude in comparison with slab gel systems, reduction in analysis time does not always translate to savings in total sample analysis time. Patient samples often must be processed in some manner before the electrophoretic separation step can be executed. Processing steps may include cell sorting, purification, DNA amplification, desalting, or labeling reactions; consequently, sample size reductions gained with the capillary format may be lost because of the material handling requirements of these initial steps. In addition to the time constraints of these steps, which are unaffected by the translation to capillaries, human intervention remains necessary between processing steps, as well as prior to the initiation of the separation step. Early on, the electronics industry realized that miniaturization in the form of the silicon microchip represented an elegant solution to the problem of complexity and has successfully exploited miniaturization to reduce the size of devices while increasing their capacity and functionality. The value in applying this same concept to the analytical sector, in particular the separations field, has been known for some time. While the efforts of Terry et al. [18] in the late 1970s to improve gas chromatography through miniaturization on a glass substrate were not fruitful, it presented the possibility that microminiaturization could do for the analytical sector what microchips did for the electronics industry. Miniaturized analytical devices, fabricated in glass using the same

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methodologies developed for the silicon microchip industry, have led to a wealth of literature that supports this concept. While glass-specific challenges exist as a result of its highly amophorous nature, the pioneering experiments of the Manz [7] Harrison [4], and Ramsey [19] groups demonstrated that, like fused silica capillaries, channels microfabricated in glass could be utilized effectively for highly reproducible separations. As a result of the channels being shorter than the typical capillaries used for electrophoresis, the field strength in a channel could be as much as 10-fold higher than in the capillary, yielding microchipbased separations that consume even less time than their capillary counterparts. For parallel processing of samples, arrays of microchannels can be etched into the glass device as simply as one channel is etched, providing a convenient solution to the parallel processing problem. However, the order-of-magnitude increase in analysis speed attainable with microchip devices over capillary-based systems highlights further the need to address the rate limiting nature of the preelectrophoresis processing steps. The true advantage of microchip devices over their capillary and slab gel counterparts stems from the fact that, in addition to channels, other structures can be fabricated into a single device. Structures that function as reaction chambers for DNA amplification or protein labeling, filters for cell sorting, or extraction/purification domains based on solid-phase media, are all conceivable. While the miniaturized size, combined with a reduction in the number of any sample handling steps, will allow for even smaller sample volumes to be utilized effectively, the benefits extend beyond sample size and reagent consumption. It has been observed that for reactions such as PCR, a decrease in size has the potential to lead to an increase in speed [20,21], reducing the time necessary for the preelectrophoresis processing steps. Creating a device with integrated functionality would begin to approach the realization of the ‘‘lab-on-a-chip’’ concept which has been discussed in the literature for years but still doesn’t exist in any concrete form. The development of a device that could accept a sample and, in an automated format, yield diagnostic results would take bold steps toward the robust ‘‘sample-in/answer-out’’ device that will be needed for the clinical sector to see value in this new technology. Such a device could function in a central laboratory environment for high-throughput analy-

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sis or in a miniaturized single-analysis mode at the site of patient care, an area in clinical medicine that continues to grow. The concept of a fully integrated microfabricated chip for clinical diagnostics will be discussed in this chapter in three sections. First, a brief description of the microchip manufacture process will be described. Second, the results of recent work using microchips as a diagnostic tool will be presented. These results will demonstrate the usefulness of this tool, simultaneously showing the need for a fully integrated device. Finally, we will present what we feel to be the important breakthroughs necessary to develop a fully functional and versatile clinical diagnostic tool. 1.2

CREATING MICROSTRUCTURES IN GLASS MICROCHIPS

The development of features in silicon or glass proceeds in a series of four steps. A metal layer is sputtered across the entire surface of the wafer followed by coating with a photoresist. The photoresist is exposed to UV radiation through a mask which was designed with the necessary features. The photoresist and metal in the feature areas are removed, and the wafer etched to produce structures in the wafer. While the idea is simple in concept, it is much more challenging in practice. The mask is an important component in this series of steps, since the mask image is transferred first to the photoresist during UV exposure and then to the metal layer. The metal layer then acts as a secondary mask for the etching of the image into the glass. The lines and sizes on the mask set the feature sizes of the final structures in the glass wafer. Metal masks are advantageous, providing resolution in the nanometer range, but are expensive and take days to produce. Filmtype masks are relatively easy to make and inexpensive, but features ⬍10 µm in size are not yet possible using current film technology [22]. Producing an integrated device requires only that the chambers, channels, and connections necessary to carry out the analytical methods of interest be contained in the original mask. Etching of the glass is carried out in a buffered solution of hydrofluoric acid. Different types of glass etch at significantly different rates, so the specifics of the etching step are determined empirically for the

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desired depth and width of the final structures. Smooth walls are typically achieved, but V-shaped channels result because as the glass is etched downward, the etch solution also acts laterally. This lateral etching often extends under the metal layer; thus, the depth of the structures in the glass are set by the amount of lateral etching which is acceptable. Features on the mask are often made smaller than they need to be to allow for this type of spreading. The etched wafer is then bonded to a second piece of glass, into which reservoirs have been drilled, to enclose the chambers and channels of the device. While most of the research on microchip devices involves glass substrates, the cost and effort necessary for current manufacture of microchips make them incompatible with the disposable-type assays that would be carried out in a clinical laboratory. There is some effort directed toward developing microchips out of polymeric-type materials. Polycarbonate, which can be injection molded, requires only that a single master mold containing the structures of interest be fabricated in metal. Microchips formed quickly and reproducibly by this process are now being investigated by ACLARA Biosciences [23]. Agilent Technologies has taken another approach, using polyimide as the material from which their microchips are constructed [24]. Polyimide is flexible, has good heat transfer properties, and is easily bonded to itself. Chambers and channels are easily formed in one piece by excimer laser ablation, which can then be bonded to a solid bottom layer and a top layer containing appropriate reservoirs. This type of technology is readily amenable to the production of three-dimensional microchips. 1.3 DIAGNOSTICALLY RELEVANT SEPARATIONS ON A MICROCHIP One of the earliest illustrations of microchip analysis with potential clinical application was a competitive immunoassay for serum cortisol demonstrated by Koutny et al. [25]. The final design of the fused silica microchip, which was used only for the separation, is presented in Figure 1A. Known amounts of antibody and labeled antigen were added to the sample of interest where the labeled antigen and the native antigen competed for antibody binding sites. Following a ‘‘pinched injection’’ using a cross channel that is slightly offset, free labeled antigen was

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Figure 1 (A) Microchip design utilized in serum cortisol immunoassay. (B) Two replicate injections of immunoassay product (IS, internal standard; Ag*, labeled cortisol; sample concentration of cortisol in water, 2.5 ⫻ 10⫺6 M. (C) Immunoassay separations of serum cortisol: 800 V/cm; 20 mM TAPS/AMPD (pH 8.8).

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then separated from an internal standard and the bound labeled antigen in the microchip. Quantifying the labeled antigen using laser-induced fluorescence allowed for determination of the amount of native antigen in the sample. The final concentration of serum cortisol could then be determined from a calibration curve. Figure 1B shows that duplicate microchip separations, when the sample was cortisol dissolved in water (not serum) with a cortisol concentration of 2.5 ⫻ 10 ⫺6 M, are fast and reproducible. Figure 1C shows three separations with different concentrations of serum cortisol. Clearly the results are reproducible and the speed sufficient to warrant clinical interest. The authors found that the protocol worked successfully in the range of 1–60 µg/dL or 30–1700 nM, which is centered on the concentrations of clinical interest. Chiem and Harrison [26] furthered this work with a microchip-based assay for the determination of monoclonal mouse IgG in mouse ascites fluid and an assay for the drug theophylline in serum samples. The important distinction here was that, although all sample preparation took place off-chip, analysis of real samples was successfully executed. Both separations could be completed in ⬍60 sec, extending microchip electrophoresis into the clinical regime based on speed of analysis. Woolley et al. [27] fabricated a microchip to perform genotyping of HLA-H, a gene with the potential for diagnosis of hereditary hemochromatosis. The authors utilized a 12-channel capillary array electrophoresis (CAE) microchip with sensitive laser-excited confocal-fluorescence detection using galvanometric scanning (based on a mirror-reflecting principle) to rapidly deliver the laser beam to all 12 channels. A restriction digestion using RsaI was performed on a PCR product of the gene segment potentially carrying the mutation. While 255-bp and 145-bp fragments are seen with the normal gene (G), the presence of the G → A transition at nucleotide 845 yields mutated fragments (A) of 255 bp and ⬃115 bp. The presence of the 115-bp fragment, therefore, provides the diagnostic basis for the genotyping. Figure 2 presents the electropherograms obtained using Mathies’s 12channel CAE microchip [27]. Deletions in an exon of the dystrophin gene that cause Duchenne/ Becker muscular dystrophy were shown to be detected by Wilding and coworkers [28] using a glass microchip. In this work, a complex series of degenerate oligonucleotide primed polymerase chain reactions

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Figure 2 Electropherograms of six HLA-H samples genotyped with twocolor fluorescent labeling in a 12-channel CAE chip.

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(DOP-PCR) and PCR were performed in glass microchips. The entire microchip was thermocycled with PCR taking 188 min to complete. Figure 3 presents the separations of the PCR product on the microchip (A-C) compared to separations using conventional capillary electrophoresis (D-F). While the microchip separation yielded a slight increase in resolution, the reduction in analysis time (less than twofold) is less than might be expected. That notwithstanding, the amplicon pat-

Figure 3 Chip separations of PCR amplicons. (A) Normal human DNA generated in a GeneAmp test tube using a conventional PCR thermocycler. (B) PCR amplicons of normal human DNA generated in a silicon/glass chip. (C) PCR amplicons of an MD-afflicted individual generated in a silicon/glass chip. (D–F) Samples as described in A–C except that product was separated using capillary electrophoresis.

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tern for a normal individual is easily distinguishable from that of a person afflicted with muscular dystrophy. Munro et al. [1] employed microchip electrophoresis for the analysis of gene rearrangement PCR products in T- and B-cells correlative with lymphoma, and compared the resolution and efficiency to that obtained with capillary electrophoresis and the gold standard, gel electrophoresis. The PCR products from normal T- or B-cells display a diverse variety of low-abundance DNA sequences, while cells from an individual positive for lymphoma will possess a predominance of a single DNA sequence. Figure 4 illustrates the analysis of T-cell lymphoma samples via the three electrophoretic techniques. The posi-

Figure 4 (A) Capillary electrophoresis of TCRG gene rearrangement products. T1 is negative/equivocal and T4 a characteristic negative sample. T2 and T3 are positive samples. (B) Chip electrophoresis of same samples.

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tive samples (T2 and T3) are clearly distinguished owing to the predominance of single DNA fragment sizes while the negative sample (T4) displays the characteristic polyclonal fragment population. The diagnosis with sample T1 was negative/equivocal owing to the conspicuous band/peak (arrow) present in the broad-banding pattern in all three electrophoretic methods. Microchip electrophoresis was also found to efficiently separate B-cell lymphoma assay PCR products, where profiles correlated well with those obtained via capillary and gel electrophoresis. Hofga¨rtner et al. [29] evaluated whether detection of a PCR product specific to the herpes simplex virus (HSV) could be carried out effectively on a microchip, and also compared the resolution and efficiency with that attainable by capillary electrophoresis and slab gel electrophoresis. It is important to detect this central nervous system infection early since it can result in encephalitis and even be fatal. Following extraction of DNA from the serum of a patient, amplification of a target sequence specific to the virus is carried out using PCR. Gel electrophoresis and Southern blot analysis reveal infection by the presence of a single amplified DNA fragment of a specific size. Extracted DNA from 33 CSF specimens provided in a single-blind manner were PCR amplified, targeting the thymidine kinase gene of HSV. Single-lane microchips were used with silanized surfaces coated with polyvinylpyrrolidone. Using the presence of the 111-bp PCR product as a signature for positive diagnosis, 100% sensitivity and selectivity were observed with the microchip when compared to conventional CE and gel electrophoresis. Figure 5 shows the microchip electropherograms for a HSV-positive control, a negative control, and a CSF specimen, with the Southern blots shown in the insets. Both this work and the work by Munro et al. [1] represent comprehensive studies comparing the translatability of the gel electrophoresis–based assays to capillary and microchip electrophoresis. 1.4 EXECUTING MULTICHANNEL SEPARATIONS ON MICROCHIPS Gel electrophoresis separations, while not rapid, easily allow for batch analysis of multiple samples in parallel. The result is often an average

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Figure 5 Chip separation (243 V/cm) of HSV PCR samples. (A) positive, (B) negative, and (C) weakly positive specimen.

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time per sample that is competitive with or better than that achieved using a serial analysis on a single-channel microchip, despite the rapid analysis time. Therefore, for microchips to effectively replace conventional gel electrophoresis methods for many (but not all) diagnostic applications, parallel sample processing will be essential. It is in this respect that the microchip format is ideal for creating analytical devices. Multiple structures can be created on a single device with no added methodological complexity or time investment than would be needed for etching a single structure. Mathies’ group has really been a force in forging this area. An example of the diagnostic power of a 12-channel microchip was seen in the previous section [27]. Their first foray involved the separation of DNA for the genotyping of HLA-H on six samples simultaneously, with a total separation time of 280 sec, leading to an average analysis time per sample of ⬃47 sec. The galvanometric laser beam scanning system developed for this was essential to the accomplishment of multichannel microchip electrophoresis (Fig. 6). The laser excited confocal-fluorescence scanning was mediated by a computer-controlled translation stage which allowed the laser to traverse the 1.1-mm-wide detection region on the chip at a rate of 1.0 cm/sec, or a periodic sweep every 0.3 sec. The resultant electropherograms from 12-channel scanning of pBR322 MspI are presented as an inset in Figure 6. The adaptation of galvanometric scanning to multiple microchannel chip electrophoresis served as springboard for the development of Mathies’ 96-channel capillary array electrophoresis system. The radial capillary array electrophoresis microplate [30] capable of simultaneous analysis of 96 samples, and the accompanying detection system utilized in this work are shown in Figure 7. Rather than rotate the entire microchip, a rotary fluorescence scanner circles in the inner portion of the microchip with positional increments at 0.05 µsec. This allows for all 96 channels to be interrogated at a position that yields the same effective separation length. Figure 8 presents the results of 96 parallel separations of pBR322 MspI samples utilizing the radial array microchip and rotary scanner. A single separation took only 140 sec, resulting in an average separation time of ⬃1.5 sec per sample. This represents a significant step in demonstrating the high-throughput capabilities of microchip electrophoresis that, until this point in time, were only conceptual.

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Figure 6 Detection system design with translational microchip stage and 12-channel separations of pBR322 MspI.

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Figure 7 (A) Ninety-six-channel radial array microchip. (B) Detection system with rotary scanning head.

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Figure 8 Ninety-six-channel separations of pBR322 MspI using a radial array microchip and rotary scanning detection system.

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An alternative to placing a microchip on a translation stage or incorporation of a moving detector system has been presented by Huang et al. [31] who developed a scanning approach that utilizes no moving parts. This approach exploits an acousto-optical deflector (AOD) for laser beam scanning. Acousto-optic deflection is based on the fact that a beam of light can be deflected if it passes through a material where acoustic waves are also present. The magnitude of deflection of the laser beam is dependent on the frequency of the acoustic waves. Since acoustic frequencies can be altered rapidly, the potential exists to have ultrafast scanning of a laser beam over a wide range of angles. Rapid changes in acoustic frequency input to an AOD can be mediated by voltages sent from a computer to a voltage-to-frequency converter (Fig. 9). Using this method, Huang et al. [31] have demonstrated multichannel scanning with DNA standard separations (Fig. 10B). The random addressability of the laser beam not only allows for self-aligned scanning but also ultrafast scanning with rates as fast as several hundred Hertz having been demonstrated. Figure 10A shows a single frame of a video signal captured during scanning at 200 Hz. 1.5 INTEGRATING FUNCTIONALITY With few exceptions, the work described thus far has focused on microchips as a platform for clinical diagnostics. However, in many respects electrophoretic separations on microchips, whether single channel or multichannel, only address part of the problem. To develop the fully integrated diagnostic platform, microchip-based preelectrophoresis sample processing must be explored. In other words, sample preparation must be performed on a scale commensurate with the separation technology; this currently does not exist. Figure 11 shows a conceptual design for an integrated genetic analysis microchip that contains independent functional domains for executing cell sorting, DNA extraction, PCR amplification, and electrophoretic analysis following direct loading of a whole blood sample. While such a design is oversimplistic and even naive as a result of the complexities associated with microto-micro (domain-to-domain) interfacing, these are the tasks that would have to be accomplished in a simple and time efficient manner for a ‘‘sample-in answer-out’’ microchip to be realized.

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Figure 9 (A) AOD scanning optical setup. 1, Beam expander; 2, lens; 4, AOD; 5, spatial filter; 6, dicroic beam splitter; 7, objective; 8, filter; 9, PMT; 10, amplifier and filter; 11, computer; 12, voltage-to-frequency converter. (B) Microchip design. (C) Closeup of detection window.

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A

Figure 10 (A) Ultrafast scanning (200 Hz) across the microchannels. (B) Separation of pBR322 Hae III digest in two microchannels.

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Figure 11 Fully integrated diagnostic device.

1.5.1

Cell Sorting

A powerful feature of a fully integrated microchip will be the ability to separate the cells of interest from other cells. For example, the removal of red blood cells (RBC) from white blood cells (WBC) so that DNA may be extracted is important, especially if PCR amplification is to be carried out downstream. A number of groups have developed techniques for sorting and moving whole cells through structures in microfabricated devices. Brody et al. [32] have developed a method for sorting blood cells. The cells travel through a microarray of various channel widths via hydrodynamic flow (channel width range from 2.5 to 4.0 µm at a depth of approximately 4 µm) Cell separations correlated not to cell size, but to the relative rigidity of the cell membrane. The authors found a significant correlation of rigidity to the presence of intracellular calcium concentration. This technique has allowed the authors to identify unusual cells from an otherwise ‘‘normal’’ population. Frazier and coworkers [33] utilized a micromachined electrical field– flow fraction system (µ-EFFF) to separate polystyrene particles of various sizes ranging from 44 to 261 nm. Particles flow through a microchannel that has a potential across it. This creates a shear force, which results in the separation based on the size of the particle. The flow profile is laminar, where the smaller particles flow faster close to the center of the parabolic flow profile and the large particles flow slowly near the wall of the microchannel. Wilding et al. [34] use a different approach to trap WBCs from whole blood using ‘‘weir-type’’ filters in silicon-glass hybrid microchips. The microchip employs a series of

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microposts, which function to trap cells. RBC navigate through the microposts, while WBC are trapped. The WBC were then used as a DNA source for microchip-based PCR, which will be discussed in more detail later. 1.5.2

Flow Control and Microreactors

Jacobson and Ramsey [35] were among the first to add additional functionality to the separation capabilities of a microchip device. This device incorporated a small volume reaction chamber where restriction enzyme-mediated digestion of DNA could be carried out. The pBR322 fragments resulting from the HinfI digestion were separated in an etched channel 67 mm in length (Fig. 12A). While the separation yielded relatively poor resolution (Fig. 12B), preelectrophoresis sample preparation was accomplished quickly with an overall analysis time (digestion through separation) of roughly 5 min. This same group furthered the concept of on-chip reaction by utilizing electrokinetic flow to mediate the mixing of substrate (resorufin β-D-galactopyranoside), enzyme (β-glactosidase), and inhibitor (phenylethyl β-D-thiogalactroside, lactose, p-hydroxymercuribenzoic acid) from different special locations on the chip [36]. The progress of the assay was monitored by detection of the hydrolysis product, resorufin, from the reaction of the enzyme with the substrate using LIF at a wavelength of 488 nm. The enzymatic assay was carried out in ⬃20 min with a fourfold reduction in volume of reactants needed in comparison with the conventional assay. With the vision that whole-cell transport would be a key feature in potential reactions on-chip, Li and Harrison [37] explored the transport of biological cells within a microchip device. They showed that whole cells could be moved through a microchannel network using electroosmotic flow or electrophoretic pumping. Figure 13 shows a photomicrograph of whole cells as they moved through the channel with flow control being accurate enough to direct all cells through the junction and into the branch maintaining a potential. The key to this work was that the authors found that cells present in the electrically floating channel remained immobile. Many reactions of biological significance are inhibited by the presence of whole cells (for exam-

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A

Figure 12 (A) Microchip design for DNA restriction fragment analysis. The deceptively long separation channel is 67 cm. (B) Electropherogram of products from digestion of pBR322 by HinfI.

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Figure 13 Flow of E. coli cells through one branch of a Y intersection.

ple, PCR is successfully inhibited in the presence of RBC). The ability to have such fine control over cell mobility will clearly be an important feature of the ‘‘sample-in answer-out’’ microfabricated device. In addition, using the same design, they were able to demonstrate the ability to manipulate and disrupt cells for analytical purposes by lysing them with sodium dodecyl sulfate (SDS). They demonstrate that, with the appropriate buffer conditions, the effective pseudo-mixing of two streams could be accomplished. While the authors did not follow lysis by a microchip-based separation, these experiments were seminal since they demonstrated that 100% of any analyte of interest can be made available for analysis via direct onchip manipulation of cells. In an attempt to use the complex voltage control necessary to perform electrophoresis coupled with other functions such as sample mixing, Jacobson and Ramsey [38] presented electrokinetically controlled parallel and serial mixing of fluids in microchip devices. Figure 14 shows the microchip design for a parallel mixer. The electrokinetic mixing is performed by simply applying a voltage of 1.0 kV to the sample and buffer reservoirs while keeping the waste grounded. This work has clear application on-chip as a means of mixing chemicals for reaction and limit of detection studies; most importantly, it illustrates

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Figure 14 Schematic of parallel electrokinetic mixing microchip. The microchip has 1.0 kV applied at buffer and sample reservoirs and is grounded at the waste reservoir. S, sample channels; B, buffer channels; and T, T-intersections.

the potential to simplify the otherwise complex electrical design of multichannel microchips by careful design of those channels. 1.5.3

Amplification

One of the most common sample preparation steps associated with clinical analysis based on genetic testing is the polymerase chain reaction (PCR). This is a powerful enzyme-mediated DNA amplification technology that allows for the detection of DNA or RNA present at very low copy numbers and underlies almost every aspect of genetic analysis. To

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execute this process on a microchip is advantageous for several reasons. The most obvious include: (1) the reduction in the reaction volume, which would yield a dramatic decrease in the cost based on the use of less Taq polymerase; (2) a more rapid and efficient amplification due to small volume temperature cycling; and (3) seamless integration of the amplification process with electrophoretic analysis. Wilding and coworkers [39,40] were the first to explore the use of silicon-glass devices as microchambers for PCR. Using a reaction volume of 10 µL in the hybrid chip and a custom-made heating device (utilizing a Peltier heater/cooler), they were able to successfully carry out amplification of λ phage DNA and C. jejuni bacterial DNA. While these experiments used volumes that were large by microchip standards and did not interface the PCR chamber with a separation domain, they provided the first intimation that PCR could be carried out in a microchip device. This group furthered their work with a microchip device that would first isolate WBC from whole blood using silicon micropost filters (described previously) [34]. PCR reaction mixture is added to the WBC and the microchamber thermocycled. As with the work that utilized just the PCR chamber, the cycle times are comparable to those achieved in a commercially available cycler. Woolley et al. [41] were the first to succeed in coupling a PCR chamber to a microchip-based separation for DNA analysis. The design used a standalone PCR device consisting of a polysilicon heater with a polypropylene insert as a PCR chamber, which is epoxied to a separation microchip (Fig. 15). The PCR reaction mixture could be rapidly heated and cooled, with transition rates of 10°C/sec heating and 2.5°C/sec cooling. The utility of this hybrid device was demonstrated with the high-speed analysis of amplified product from the βglobin gene in Salmonella DNA. The PCR amplification was performed in 15 min (30 cycles, 96°C for 2 sec, 55°C for 5 sec, and 72°C for 2 sec), with separation times of approximately 100 sec. This study provided the first vista of how integrated PCR/electrophoretic analysis could speed genetic analysis. Kopp et al. [42] defined a clever approach to thermocycling using continuous flow on a microchip. This involved mediating PCR amplification of DNA by hydrostatically pumping the PCR reaction mixture through a serpentine channel that passed through three discrete temper-

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Figure 15 (A) Schematic of PCR-CE integrated device, including detection system design. (B) Closeup of PCR chamber. (C) Junction between polypropylene insert and microchip.

ature zones on the microchip (Fig. 16). The amount of PCR product observed can be controlled by the number of cycles included on the microchip or the rate at which the PCR mixture flows through the microchannel. In this work the 176-bp PCR product was collected and detected off-chip via gel electrophoresis. However, there are some disadvantages associated with this design. One problematic area is the flow control. Sample is hydrostati-

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Figure 16 Design of continuous-flow PCR microchip. (A) A microchip for flow controlled PCR. (B) Layout for continuous-flow PCR microchip.

cally pumped through the temperature regions. This adds another level of complexity to a fully integrated chip, which has relied mainly on electrostatic driven fluid motion. This mechanism of PCR limited the flexibility of the reaction; the number of cycles is constant, which implies that a single chip has a truly limited functionality. Lastly, and perhaps most significant is the size of the microchips utilized in this study. The microchips are rather large, approximately 40 ⫻ 40 mm; in a field where the goal is to pack as much chemistry in as small an area as possible, the size of the continuous flow system is too great. Work by Waters et al. [43] represents one of the first real examples of a higher-ordered integrated device. The microchip device served as a multifunctional platform for the chemistries associated with cell lysis, polymerase chain reaction, and electrophoretic separation. E. coli cells were loaded into a reservoir on the microchip. While not very practical, the entire microchip was thermocycled in order to lyse the cells and then amplify select DNA targets via the polymerase chain

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reaction. A sample from the reaction chamber was transported into the electrophoresis channel using electro-osmotic flow, and separation was carried out over a sieving gel. While E. coli DNA, specifically a cDNA clone of the plant gene agNt84, was successfully amplified, the thermal mass constraints associated with having to temperature-cycle the entire microchip in a conventional thermocycler, led to cycle times that were long. Consequently, an advantage associated with the ability to rapidly detect the amplified DNA was offset by the 33/4 hr required for lysis and amplification. A recent presentation of work from the Mathies group detailed a microfluidic DNA device containing an integrated chamber for PCR, interfaced with an electrophoresis domain [44]. Utilizing microfluidic valves and vents, they were able to control sample loading into the 280-nL reaction chamber. A thin-film resistive heater regulated temperature in the PCR chamber, resulting in cycles that were 30 sec in length. After 20 cycles, they were able to effectively amplify and detect a 136bp amplification product of the M13/pUC19 cloning vector from five to six initial copies. Burns et al. [45] have approached the development of an integrated microchip device from an engineering-oriented perspective and, consequently have developed a much more sophisticated, yet functional, device for amplification and analysis. Incorporated into their microchip design is a region for PCR thermocycling utilizing resistive heaters and temperature sensors, a region for sample loading, and a channel filled with a gel matrix for sieving-based separations. The DNA analysis device was used to successfully amplify a 106-bp fragment of DNA in a strand displacement amplification experiment. Their work presents the first rapid nanoliter volume DNA amplification directly coupled to a separation channel using the microchip format. This work by Ramsey and Burns represent important steps forward in developing the fully integrated microchip, particularly smallvolume PCR on the same device as the separation channel. These studies clearly present results that will catalyze the transformation of PCR on microchips from its current infantile stage to a substantiated miniaturized sample preparation technique. The kinetics for extending, annealing, and denaturing DNA in PCR are fast in comparison to the time scale of a typical temperature cycle. The major limitation is the thermal

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load of traditional PCR and the Peltier heating/cooling technology. While a single cycle using existing technologies takes on the order of 3 min, it is not unreasonable to expect that the cycle time could be reduced to 10s of seconds with the possibility of fewer cycles needed if the amplification in small volumes is more efficient. It is interesting that all of the approaches described thus far involve some form of ‘‘contact’’ heating, of the entire chip [43], a portion of the chip [39–41], or through use of resistive heating within the microchamber itself [44,45]. Novel approaches that do not mimic ‘‘macro’’ methods for PCR have been described and provide additional avenues to explore. Both Northrup [46] and Oda et al. [21] have described alternative concepts for PCR amplification based on ‘‘noncontact’’ thermocycling using infrared (IR) heat. This approach may be advantageous for several reasons. First and foremost is thermal load. With the exception of the resistive heating described above, most approaches involve heating the PCR mixture by heating its environment. Thermal mass constraints limit the rate at which the temperature can rise and fall; cycle times can become relatively long [43]. As a result of the fact that IR light can selectively excite a vibrational band of water (and not the substrate in which it is contained), IR heating limits concerns of thermal load to only the volume of the PCR mixture to be temperature cycled. Logic dictates that by reducing the volume of water, the thermal load is reduced and, hence, faster IR-mediated thermocycling can be carried out. Hu¨hmer and Landers [47] have shown that nanoliter volumes of PCR mixture can be thermocycled in a capillary with average cycle times on the order of 3 sec. The work carried out using capillaries has been extended to the microchip format. Figure 17 illustrates the design of the IR heating system. A dummy chamber with a thermocouple is used to regulate the power to the tungsten lamp, which is controlled by a computer using Labview. IR-mediated thermocycling has been applied to glass, glass/polydimethylsiloxane hybrid, and polyimide microchips. Thermocycling of the polyimide microchip is presented in Figure 18. A single cycle takes only 17 sec. With rapid thermocycling comes a reduction in reaction volume. This is advantageous because it allows more seamless integration of a PCR chamber with a separation channel. Unfortunately, the reduction in volume comes with at least a 1000-

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Figure 17 Design of noncontact infrared light source for PCR. (1) Computer utilizing Labview for lamp control and temperature measurement; (2) type T thermocouple; (3) gold mirror; (4) microchip with dummy chamber; (5) 695-nm longpass filter; (6) lens for lamp focusing; and (7) 50-W tungsten projector bulb.

fold increase in the surface-to-volume ratio of the PCR chamber when compared to conventional polypropylene reaction tubes for commercial thermocyclers. This results in the deactivation of Taq polymerase via adsorption to the surface of the PCR chambers. Many solutions have been presented in the literature ranging from addition of bovine serum albumin, for nonspecific binding to the chamber surface to maintain more enzyme in solution, to coatings for the PCR chamber similar to those employed in separation chambers. 1.5.4

Purification/Extraction

Sample preparation in clinical diagnostics is generally time-consuming. Seamless integration of DNA purification with DNA amplification and

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Figure 18 Thermocycling profile in a polyimide microchip. Total cycle time is 17 sec.

DNA detection into a single microfabricated device therefore holds great potential in clinical diagnostics. For molecular diagnostics, there are a variety of DNA purification methods to prepare PCR-suitable DNA; the most common methods are organic solvent extraction, ultracentrifugation, and ion exchange chromatography. Most of the commonly utilized DNA purification methods are either not compatible with the microchip format or not easily integrated. It is well established that DNA can be absorbed by silica, and since the most common substrate for microchips is silica, it seems logical that silica-based adsorption of DNA should be explored for microliter and submicroliter scale DNA extraction. Tian et al. have taken the first steps in evaluating this [48]. They show that conditions can be defined for purifying human genomic DNA directly from human white blood cells, and even from whole blood, by selective adsorption onto a ⬃500-nL bed of silica particles in a capillary. Using fluorescence assays for both protein and DNA, they found about 70% DNA could

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be recovered and ⬎80% of the protein removed from a few microliters of blood by micro-solid-phase extraction (µSPE). The capacity was in the range of 10–30 ng/mg silica particles, and extraction could be carried out in ⬍10 min. They showed the suitability of silica-based microextracted DNA from white blood cells, cultured cells, and whole blood for successful PCR amplification of BCRA1 and BRCA2 mutations. This time frame is compatible with the analysis time of microfabricated chip electrophoresis. Efforts of this group are focused on integrating this methodology into the microchip platform. 1.6

CONCLUDING REMARKS

It is our hope that we have been able to present some of the more interesting forays in the realm of the integrated diagnostic platform. This is by no means a complete recounting of the many papers that have contributed to this area. In a growing field, this is just the beginning of work that will afford rapid clinical diagnostics. REFERENCES 1. Munro NJ, Snow K, Kant JA, Landers JP. Clin. Chem. 1999, 45, 1906– 1917. 2. Rossier JS, Schwarz A, Reymond F, Ferrigno R, Bianchi F, Girault HH. Electrophoresis. 1999, 20, 727–731. 3. Liu S, Shi Y, Ja WW, Mathies RA. Anal. Chem. 1999, 71, 566–573. 4. Chiem NH, Harrison DJ. Electrophoresis. 1998, 19, 3040–3044. 5. Walker PA, Morris MD, Burns MA, Johnson BN. Anal. Chem. 1998, 70, 3766–3769. 6. Woolley AT, Lao K, Glazer AN, Mathies RA. Anal. Chem. 1998, 70, 684–688. 7. von Heeren F, Verpoorte E, Manz A, Thormann W. Anal Chem. 1996, 68, 2044–2053. 8. Jacobson SC, Ramsey JM. Electrophoresis. 1995, 16, 481–486. 9. Deforce DL, Millecamps RE, Van Hoofstat D, Van den Eeckhout EG. J. Chromatogr. A. 1998, 806, 149–155. 10. Siles BA, Collier GB, Reeder DJ, May WE. Appl. Theor. Electrophor. 1996, 6, 15–22. 11. Guttman A, Starr C. Electrophoresis. 1995, 16, 993–997.

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12. Pancholi P, Oda RP, Mitchell PS, Landers JP, Persing DH. Mol. Diagn. 1997, 2, 27–37. 13. Mendoza LG, McQuary P, Mongan A, Gangadharan R, Brignac S, Eggers M. Biotechniques. 1999, 27, 778–788. 14. Gao Q, Pang HM, Yeung ES. Electrophoresis. 1999, 20, 1518–1526. 15. Xue G, Pang HM, Yeung ES. Anal. Chem. 1999, 71, 2642–2649. 16. Tan H, Yeung ES. Anal. Chem. 1998, 70, 4044–4053. 17. Simpson PC, Roach D, Woolley AT, Thorsen T, Johnston R, Sensabaugh GF, Mathies RA. Proc. Natl. Acad. Sci. USA. 1998, 95, 2256–2261. 18. Terry SC, Jerman JH, Angell JB. IEEE Trans Electron Devices. 1979, 26, 1880–1886. 19. Waters LC, Jacobson SC, Kroutchinina N, Khandurina J, Foote RS, Ramsey JM. Anal. Chem. 1998, 70, 5172–5176. 20. Hu¨hmer AFR, Landers JP. Anal. Chem. 2000, 72, 5507–5512. 21. Oda RP, Strausbauch MA, Hu¨hmer AFR, Borson N, Jurrens SR, Craighead J, Wettstein PJ, Eckloff B, Kline B, Landers JP. Anal. Chem. 1998, 70, 4361–4368. 22. Duffy DC, McDonald JC, Schueller OJA, Whiteside GM. Anal. Chem. 1998, 70, 4974–4984. 23. McCormick RM, Nelson RJ, Alonso-Amingo MG, Benvegnu DJ, Hooper HH. Anal. Chem. 1997, 69, 2626–2630. 24. Trout AL, Hu¨hmer AFR, DiMartini A, Lang T, Swedberg S, Udiara S, Landers JP. Presentation, HPCE, 1999. 25. Koutny LB, Schmalzing D, Taylor TA, Fuchs M. Anal. Chem. 1996, 68, 18–22. 26. Chiem N, Harrison DJ. Anal. Chem. 1997, 69, 373–378. 27. Woolley AT, Sensabaugh GF, Mathies RA. Anal. Chem. 1997, 69, 2181–2186. 28. Cheng J, Waters LC, Fortina P, Hvichia G, Jacobson SC, Ramsey JM, Kricka LJ, Wilding P. Anal. Biochem. 1998, 257, 101–106. 29. Hofga¨rtner WT, Hu¨hmer AFR, Landers JP, Kant JA. Clin. Chem. 1999, 45, 2120–2128. 30. Shi Y, Simpson PC, Scherer JR, Wexler D, Skibola C, Smith MT, Mathies RA. Anal. Chem. ASAP article. 31. Huang Z, Munro N, Hu¨hmer AFR, Landers JP. Anal. Chem. ASAP article. 32. Brody JP, Han Y, Austin RH, Bitensky M. Biophys. J. 1995, 68, 2224– 2232. 33. Gale BK, Caldwell KD, Frazier AB. IEEE Trans. Biomed. Eng. 1998, 45, 1459–1469.

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34. Wilding P, Kricka LJ, Cheng J, Hvichia G, Shoffner MA, Fortina P. Anal. Biochem. 1998, 257, 95–100. 35. Jacobson SC, Ramsey JM. Anal. Chem. 1996, 68, 720–723. 36. Hadd AG, Raymond DE, Halliwell JW, Jacobson SC, Ramsey JM. Anal. Chem. 1997, 69, 3407–3412. 37. Li PCH, Harrison DJ. Anal. Chem. 1997, 69, 1564–1568. 38. Jacobson SC, McKnight TE, Ramsey JM. Anal. Chem. 1999, 71, 4455– 4459. 39. Cheng J, Shoffner MA, Hvichia GE, Kricka LJ, Wilding P. Nucleic Acids Res. 1996, 24, 380–385. 40. Shoffner MA, Cheng J, Hvichia GE, Kricka LJ, Wilding P. Nucleic Acids Res. 1996, 24, 375–379. 41. Woolley AT, Hadley D, Landre P, deMello AJ, Mathies RA, Northrup MA. Anal. Chem. 1996, 68, 4081–4086. 42. Kopp MU, deMello AJ, Manz A. Science. 1998, 280, 1046–1048. 43. Waters LC, Jacobson SC, Kroutchinina N, Khandurina J, Foote RS, Ramsey JM. Anal. Chem. 1998, 70, 158–162. 44. Lagally ET, Simpson PC, Mathies RA. Presentation, Frederick Conference on Capillary Electrophoresis, 1999. 45. Burns MA, Johnson BN, Brahmasandra SN, Handique K, Webster JR, Krishnan M, Sammarco TS, Man PM, Jones D, Heldsinger D, Mastrangelo, CH, Burke DT. Science. 1998, 282, 484–487. 46. Northrup PCT. Patent. 48. Tian H, Hu¨hmer AFR, Landers JP. Anal. Biochem. 2000, 283, 175–191.

2 Creating a Lab-on-a-Chip with Microfluidic Technologies Anne R. Kopf-Sill, Andrea W. Chow, Luc Bousse, and Claudia B. Cohen Caliper Technologies Corp., Mountain View, California

2.1 INTRODUCTION Miniaturization and integration of electronic circuits has both enabled and driven the development of the information industry over the past five decades. The hope is that these same two features will be applied to the biotechnology industry to revolutionize the speed and power of biochemical and biological assays thus creating an explosion of information. In the case of electronic circuits, the shrinking feature size generates faster calculations owing to the better frequency response of smaller devices. Calculations are further accelerated because the smaller size accommodates more devices and functions on a single chip, reducing the number of slow transitions to the outside world. Laboratory protocols usually involve combinations of reactions, separations, and detection steps—often repetitively. Making the analogy to the electronic circuits, the separation steps are inherently faster at smaller-length scales. However, unlike electronic circuits, biochemical reactions and product detection steps are not inherently faster at small-length scales. Yet, these reactions can often be driven faster by increasing the concentrations of reagents to very high levels. Pushing 35

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the reaction by increasing the concentration of reactants is feasible because microfluidic devices use nanoliter volumes. The speeds of these reactions and analyses are further enhanced by the exquisite control microfluidics uses for mixing, reaction times, and temperatures. The superior control of these parameters permits the reliable use of kinetic measurements, avoiding the time of waiting for reaction completion. This review article describes microfabricated devices used in microfluidic technologies and examines the modes of fluid actuation and mechanics of these movements. This is followed by a review of the progress made to date on the integration of analytical systems on labon-a-chip devices. Integration refers to the combination of two or more procedures onto one fluidic device. The procedures could include dilution of sample or reagents, mixing of two or more reagents, separation of reactants or products, heating of reactants such as in the PCR reaction, and detection. Of course, the integration is most valuable when the multiple steps are performed automatically, without intervention. 2.2

DESCRIPTION OF DEVICES

Lab-on-a-chip devices are typically made of two pieces of glass, quartz, silicon, or plastic. In the case of the first three substrates, the fluid channels are photolithographically etched into one of the pieces [1]. The second layer is bonded, usually thermally, to the etched layer. It is crucial to perform the bonding in a clean room as the key step is removal of all particulates. Early chips were made of silicon but this substrate has largely been abandoned in favor of glass, which can tolerate the electric fields required for electrophoretic separations. Many researchers have made microfluidic devices from plastic substrates instead of glass or silicon with the hope that these devices will be cheaper to mass produce. The fabrication techniques include laser ablation [2], thin-film lamination [3], and embossing with a wire or etched silicon master [4]. The embossing technique has been used for PMMA (polymethylmethacrylate) substrates; flexible silicone rubber (polydimethylsiloxane) has been cast into chip devices [5]. Bonding plastic devices is a challenge because the cross-sectional dimensions must be maintained and any gaps between the top layer and channel layer must not permit measurable amounts of fluid or sample to enter.

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One way to avoid this problem is to use an adhesive layer as the top surface. However, this method produces a channel with different electro-osmotic properties in the channel surfaces, which can cause dispersion in sample plugs. After the channels have been etched, holes are created in one of the layers to enable the user to load samples or reagents into the channels. These holes are made with conventional machining methods. The major advantage of a lab-on-a-chip device is flexibility; you can rapidly reconfigure the chip design for a specific type of assay and you can program the timing and dilution precisely, at the time of the assay. In most cases, the masks used to define the fluidic channels in a chip are inexpensive and easy to create. This gives the fluid designer a wide range of control over the complexity and design of the fluid network Figure 1 shows a picture of a typical glass microchip device.

Figure 1 A glass microfluidic chip from Caliper. The channels, filled with dye in this picture, are created photolithographically and end in circular reservoirs that are filled with reagents or test compounds.

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MICROFLUIDICS Fluid Movement

Several different modes of fluid actuation are used on microfluidic chip devices: electrokinetics, pressure, electrohydrodynamics, capillary pressure, centrifugation, and surface tension gradients. Electrokinetic movement is the most studied and widely used motive force in integrated assays. Electrokinetics Electrokinetics refers to the linear combination of electro-osmosis and electrophoresis. Electro-osmosis drives bulk-fluid movement and electrophoresis drives the movement of chemical species on the basis of their charge. Electrokinetics has the advantage of speed and accuracy; there are very rapid fluid switching and very rapid separation of chemical species with a charge-to-mass difference, both of which are controlled by a computer. The fluid movement is not affected by the dimensions of the channels and so can be configured at assay time by changing the electric filed in the channel. In electro-osmosis, the most important property of the flow is a flat velocity profile. This means the dispersion encountered by a finite plug is very small. Electrokinetics has the disadvantages of being sensitive to pH, ionic strength of the buffer, and the surface activity of the compounds being tested. This limits or dictates the choice of buffers and additives in an assay. There are two basic injection formats for microfluidic devices using electrokinetic actuation: cross injection (also called plug injection) and gated injection. In cross injection, the dye or reagent is loaded from one side of a channel straight across and then injected into the cross channel (Fig. 2, top panel). The dimension of the area intersected by the two channels primarily determines the amount injected. In this format a very narrow and well-defined plug is introduced. This injection mode is useful when excellent resolution is required. In gated injection, the dye or reagent is introduced around a corner and the fluid is injected by applying the voltage through the intersection for a variable length of time. The amount injected is determined by the time and field strength of the injection (Fig. 2, bottom panel). Besides allowing vari-

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Figure 2 Microfluidic injection schemes for introducing finite plugs of reagents into channels. The top panel is a schematic of cross injection. Reagent is first loaded straight across followed by injection into the cross channel. The bottom panel shows gated injection. Reagent is loaded around a corner and then fluid is injected by applying a pulsating voltage.

able injection amounts, the quantity of reagent introduced into the channel can be much larger in the gated injection than in the cross injection scheme. A variation of the cross injection is the offset cross injection, or double-T [1], as Jed Harrison calls it [6]. In this case, the inlet and outlet are offset from each other and much larger injection volumes can be injected.

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It is useful to be aware of the two domains in which one can analyze moving reagents in microfluidic devices because they have interesting implications for the injection scheme. One is the space domain; the other, the time domain (Fig. 3). Space domain is the view resulting from a photograph taken of the channel—therefore an instant in time. If reagent 1 is equal in volume to reagent 2, that would mean the two plugs were the same length in the recorded photograph (Fig. 3, top panel). If analyzed in the time domain using a PMT or other point detector data recorder, from a fixed position over time, two reagent plugs that are the same length in the channel but that move at different velocities will record as peaks or plateaus of different time lengths (Fig. 3, bottom panel). In the cross injection, equal volumes of multiple species are injected and exhibit the same size plug in the space domain. As charged chemical species with different mobilities move down a channel, their migration varies; this is time domain. In the gated injection, the species plug lengths vary in the space domain but are equal size in the time domain.

Figure 3 Schematics of space and time domains.

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Pressure Vacuum or pressure, usually hydrostatic head, is used to perform certain functions on microfluidic devices. Pressure movement has the advantages of being insensitive to chemical properties of the buffers or surface reactivity of the reagents. The disadvantage, however, is that the fluid movement is not constant across the channel, thereby causing hydrodynamic dispersion. In systems with a single pressure controller as in the Kopp et al. [7] example described below, the chips must be designed to perform specific dilutions and assays. One of the most interesting lab-on-a-chip assays uses pressure to demonstrate a continuous chemical reactor. Using hydrostatic pressure as the fluid actuator, Kopp et al. [7] performed rapid PCR cycles on nanoliter fluid volumes. Wilding and Kricka [8] also used pressure to prepare blood for PCR reactions. They used a syringe pump to move blood over a wire, capture the white cells, rinse and flush the cells with PCR buffer, and then cycle the temperature of the chip on a Peltier heater-cooler device. Researchers at Orchid Biocomputer used a pressure driven pump to perform DNA hybridization on beads in a microchannel held in place with a magnetic field [9]. Burns et al. [10] have recently demonstrated an integrated DNA analysis assay using pressure and electrokinetic modes of reagent movement. To date no one has used only pressure to perform multiple types of analyses on a microfluidic device. Surface Tension and Capillary Action Other modes of fluid movement on microfluidic devices include surface tension gradients and capillary action. In addition to the work mentioned above integrating pressure and electrophoresis, Burns et al. [11] have made other integrated devices that use the change in surface tension with temperature to move nanoliter-size fluid drops on a chip. Researchers at Orchid Biocomputer used capillary forces in addition to pressure pulses to control fluid movement in microfluidic devices [12,13]. Investigators at Gamera Biosciences used passive valves based on capillary action plus centrifugation to regulate fluid movement in microfluidic devices [14]. These are successors to fluidic devices developed at Biotrack and Abaxis. Although these earlier fluidic devices (made by injection-molding techniques) used capillary and cen-

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trifugal forces to prepare blood samples and mix reagents, they were too large to be considered microfabrication [15–18]. Combined Electrokinetic and Pressure Electrokinetic movement can have the following disadvantages: compound mixtures separate into components; test compounds move at a different speeds, dependent upon their electrophoretic mobility; and highly charged molecules may not migrate at all. Researchers at Caliper have developed a special method for electroosmotic pumping that uses two buffers with two ionic strengths. The compounds are placed in the relatively high-salt buffer and spaced by making pulsed injections with a low-salt buffer. The electric field is then dropped across the spacer buffer but not the sample-containing buffer. In this way the electrophoretic migration is decoupled from electro-osmotic flow and forces all chemical species to travel with the same velocity. This is demonstrated by experiments in which cross injections were made with a mixture of two dyes in a high-salt solution of 100 mM borate buffer at pH 8.9 (conductivity ⫽ 2.1 mS/cm). The low-conductivity buffer was 20 mM borate, also at pH 8.9 (conductivity ⫽ 0.54 mS/cm). In the standard system with one buffer, the two fluorescent dyes fluoroscein and rhodamine separate rapidly, as shown in the top panel of Figure 4. The same dyes contained in the high-conductivity buffer and surrounded by lowconductivity buffer stay together as shown in the bottom panel of Figure 4. In this system the fluid motion is a combination of electrokinetic flow in the low-salt buffer and pressure-driven flow in the high-salt buffer. The velocity profile has features of both the flat electrokinetic profile and the parabolic pressure-driven profile. The fluid mechanics is complex in that the fluid movement is driven by the salt concentration distribution and that distribution diffuses and disperses in response to the fluid motion. The use of numerical tools has helped to understand the details of this mixed-mode flow [19]. 2.3.2

Mixing

Mixing is an integral part of the microchannel design in lab-on-a-chip devices. In the everyday world, turbulent motion of fluids is used to

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Figure 4 Electrokinetic separation and containment of two electrically disparate dyes. The top panel shows repeated injections of fluorescein and rhodamine dyes in a single buffer. The bottom panel shows the same two dyes comigrating when they are dissolved in a high-ionic-strength buffer and driven with a low-ionic-strength buffer.

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mix substances. However, turbulence is only realized in systems in which inertia dominates viscous forces, and this is usually not true in microdevices. The dimensionless group, Reynolds number, is a measure of inertial to viscous forces. It is defined as Re ⫽ ρUd/η, where ρ is density, U is linear velocity, d is the cross-sectional dimension, and η is viscosity. To achieve turbulent flow one typically must have Re ⬎2000 in a circular pipe, for instance. In microfluidic devices transporting liquids (not gases), Re is ⬍1 and often ⬍0.01. Typical numbers for these fluid properties on microchips are 1.0 g/cm 3, 0.1 cm/sec, 0.001 cm, and 0.01 g/cm-sec for the variables ρ, U, d, η. Molecular diffusion is the phenomenon used for mixing on microdevices. In the macro world, diffusion is a very slow process for mixing, but in microfluidic devices the dimensions are very small and so diffusion is very effective at mixing. A rule of thumb for mixing time is: t ⫽ x 2 /2D. For small molecules, a typical diffusion coefficient, D, is 3 ⫻ 10 ⫺6 cm 2 /sec so the time for mixing of two fluid streams in a channel 50 µm (0.005 cm) wide is about 4 sec. At flow rates of 1 mm/sec this requires a mixing channel length of 4 mm which is easily designed into the fluid circuit. Of course, for larger molecules such as proteins, the time for diffusional mixing is much longer. Luckily, one does not need all reactants to be uniform across the channel to yield quantitative data. Most reactions of interest are first order (i.e., the rate is linearly proportional to the concentration of each reactant). For these reactions, only one of the reagents needs to be uniform across the channel. The variable concentration of the nonuniform reagent produces variable product across the channel but the average concentration is as expected for a uniform reagent. Reactions between a large and small molecule, such as an enzyme and substrate, still only require ⬃4 sec of mixing time. The reaction between two large molecules, an uncommon but not impossible case, would require a longer mixing time; for diffusion coefficients of 5 ⫻ 10 ⫺7 cm 2 /sec, the time is 25 sec and for velocities of 1 mm/sec, 25 mm of mixing channel length is required. 2.3.3

Chip Length Scales

Microfluidic devices have channel widths and depths in the 10- to 100µm range. In channels larger than this, mixing becomes problematic.

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In channels smaller than this, the detection step becomes difficult. A 1-mm channel length has about 1 nL of volume, in a 10 ⫻ 100 µm channel. Some assays require concentrations of biological molecules in the 1-nM range. This is equivalent to one femtomole of material or only 10,000 molecules in this 1 mm of channel length. Dimensions 10fold smaller would still provide sufficient molecules for a statistically meaningful result, but lower numbers would make detection unreliable. Thus the optimum channel size is around 10 ⫻ 100 µm. The trajectory of microfluidic devices does not follow the same size reduction path as electronic devices. For lab-on-a-chip applications there is a finite number of molecules required for detection [20]. 2.3.4

Conservation of Flux

A fascinating phenomenon in electrokinetically driven chips is the relationship between velocity and concentration. In test tube experiments or pressure-driven flow systems, the concentration of product is independent of the mobilities of the reactants and products. In electrokinetic systems this is not true—the mobilities of various components can affect the product concentrations produced on the chip. The product of a reaction frequently has a different charge than the original molecule. If two fast reactant molecules combine to make a slow product molecule, the concentration of product will be higher on an electrokinetically driven chip than in the equivalent test tube or standard-flow system experiment. In the chip experiment this phenomenon may result in a more sensitive measurement. To enhance the concentration of a product, this phenomenon can be exploited by tuning a system to match the electro-osmotic flow with the electrophoretic mobility. It should be noted, however, that the inverse result is also possible. If a faster product is produced, detection can be more difficult. 2.4 ASSAYS 2.4.1

Enzyme Assays

There are many examples in which researchers have demonstrated kinetic parameters of enzymatic reactions on chips. Most of these examples used fluorescence detection. Ramsey et al. [20] were the first to

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use a chip for an integrated enzyme assay with the measurement of βgalactosidase [21]. Using a single reagent port to vary the substrate concentration, they were able to obtain accurate Michaelis-Menten kinetic data. Measuring both protease and phosphatase enzymes, Cohen et al. [22] went one step further by having multiple reagent ports which controlled the concentration of each component (Fig. 5). This allowed total on-chip control of all concentrations in an enzymatic reaction. Again, reproducible kinetic data were easily obtained using microfluidics. The monitoring of kinase activity with microfluidics is the most exciting—primarily because these enzymes are extremely important in biological systems but traditionally difficult to assay. Researchers at Caliper have successfully developed a microfluidic system that integrates all the steps of a kinase reaction as well as the electrophoretic separation of the phosphorylated product [23]. This was achieved by using microchips that were designed with both gated and cross injectors as shown in Figure 6. The upper looped portion of the chip (along with

Figure 5 Chip design allowing simultaneous control of concentration of three reagents.

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Figure 6 Chip and assay designs and for a nonfluorogenic enzyme assay using both cross and gated injection schemes.

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(A)

(B)

Figure 7 Microchip kinase assay data. The panel on the left shows fluorescent peaks of substrate, dye marker, and product. Right-hand panel is K m determination.

the flow rate) dictates the reaction time. The resultant mixture is injected into the lower looped portion where the substrate and product are separated. Figure 7A shows a sample of the separation data and analysis. The three peaks are the substrate, a neutral marker, and the product. The K m is evaluated in a double-reciprocal plot of the rate and substrate concentration (Fig. 7B). The equivalent experiment in a macro format requires timing and then stopping the reaction and running the product on a separation device. Here the chip does the timing, stopping, and separation automatically. The real promise for measuring kinase enzyme activity with a microchip is that it enables fast, flexible assay formats that require very little reagent. 2.4.2

Immunoassays

Clinical diagnostics is the main application for immunoassays on microdevices. The challenges in working with the variability in patient

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samples and long incubation times required for some immunoassays have limited the development of these systems. Several groups have demonstrated microchip immunoassays for cortisol, thyroxine, and theophylline [24–26]. In these examples, the incubation of analyte and immunoreagent was done off the chip and the separation step performed on the chip. In 1997 Chiem and Harrison [25] performed the first integrated immunoassay that included mixing, incubation, and separation on one microfluidic device. 2.4.3

Cell-Based Assays

Li and Harrison [27] developed one of the first cell-based assays that used a fluidic device. Cells were pumped across the device together with a reagent that caused cell lysis. Wilding and Kricka [8] used whole blood as the starting DNA material on a PCR microfluidic device in which white cells were captured on a shallow shelf of the device. Very recently, a group at Caltech demonstrated cell sorting on an electrokinetic microfluidic device [28]. Like a conventional cell sorter, it detects a fluorescent signal and diverts the beads or cells to a distinct well. However, unlike the conventional FACS, it is capable of ‘‘reverse’’ sorting. In this case, when a cell of interest is detected, the flow is reversed and pumped slowly enough to switch the stream and capture the desired cell. This technique is especially useful in rare cell collection as it decouples the speed of detection from the speed of diversion. 2.4.4

PCR

One of the most interesting and demanding applications for integrated analysis is the PCR reaction. A simple system to perform PCR and electrophoretic separations on a single chip has the PCR reagents in a fluid reservoir, thermally cycles the entire chip, and injects the product from the well for analysis [29,30]. The advantages of using microfabricated devices are speed, convenience, and sample size reduction. The smaller dimensions enable faster thermal cycling because the thermal mass is tiny; integrated heating devices further decrease the cycling time because of efficient thermal contact. The integration of sample preparation, amplification, and analysis on a single microdevice provides an easy-to-use assay that

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requires less sample material because there are no off-the-chip analysis steps. In 1994 Wilding et al. [31] first demonstrated PCR on a microfluidic device made of glass-silicon. They performed an integrated assay that included a cell-sample preparation coupled to PCR [8]. They created a chip with barriers that allowed the red cells cells that inhibited the PCR assay to pass but trapped the white cells. Researchers at Lawrence Livermore and Berkeley successfully combined amplification and analysis on a single device [32]. The device was composed of two pieces: a silicon sandwich for thermocycling and a glass sandwich for separations, assembled together with epoxy. The cycle time was only 30 sec on a PCR sample of 20 µL and the integrated assay including detection was only 20 min. Several hours would be required to achieve the same information in conventional equipment. Several other systems have been developed which monitor the reaction in real time. This method enables one to quantify the starting DNA and to stop an amplification at the point where subsequent analyses are the most sensitive. The most common systems use either intercalating dyes that monitor the production of double-stranded DNA or Taqman probes which monitor specific sequences of DNA. Northrup has recently reported a cycle time of 17 sec and a total PCR amplification time of 7 min [33]. There are two examples of submicroliter PCR. One of these, produced by Andreas Manz and his coworkers, is a novel system in which fluid channels meandered across hot blocks at set temperatures [7]. The advantage is a very rapid cycle time of only 4.5 sec and a total time of 90 sec for 20 cycles. The disadvantage of this system is that the number of cycles is predetermined and the efficiency is low, requiring a high starting concentration of material. In this configuration, the detection was off-chip, so each sample had to be run long enough to produce microliter quantities of material for detection off-chip. Another system, demonstrated by Kalinina and coworkers [34] amplified and detected 10 nL of material using the Taqman assay described above. Capillaries were placed on heating blocks and thermally cycled with hot and cold air. This format is not amenable to integration. When the capability to amplify and detect at a nanoliter scale is developed, it will enable true integration of PCR.

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Figure 8 Microchip with attached capillary for high-throughput sample accession.

2.5 SAMPLE AND COMPOUND ACCESSION One of the most important issues for high-throughput applications for lab-on-a-chip devices is sample acquisition. The sample capacity for analysis on a chip is high but limited by the speed with which samples can be brought to the chip. Researchers at Caliper use a capillary tube attached to the chips to bring 1- to 5-nL samples onto the chip for testing with sample spacing of 5–20 sec (Fig. 8). For higher throughput, multiple capillaries attached to one chip feed samples even faster. 2.6 MICROFLUIDIC VERSUS ARRAY TECHNOLOGIES One of the features that distinguishes microfluidic devices from arraybased technology is the serial versus parallel nature of the formats. This difference has profound effects on the calibration scheme used to produce reliable results. Any variation in a system (e.g., lamp fluctua-

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tions, reagent degradation) that has a time scale of minutes or longer can be calibrated out by running control or test samples. These can be inserted in a sample stream as often as needed. The ability to ensure quality results is thus much higher than for single-use devices, where the device is spent during testing.

2.7

CONCLUSIONS

Microfluidic devices offer a flexible format to perform a wide variety of assays. Many different laboratory functions have been demonstrated on chips by researchers in academia and industry. These include enzyme assays, PCR, blood analysis, and cell-based assays. The modes of fluid movement for these demonstrations include electrokinetic, pressure, capillary action, and surface tension gradients. The first examples of integration of multiple functions are now being realized and the difficulties addressed. The pace of microfluidic experimentation is expanding rapidly, and we can expect the promise of miniaturization and integration in the biochemical laboratory to become apparent over the next few years.

REFERENCES 1. Effenhauser, C. S., Manz, A., et al. (1993). Glass chips for high-speed capillary electrophoresis separations with submicrometer plate heights. Anal. Chem. 65: 2837–2842. 2. Robert Elghanian, J. (1997). Selective colorimetric detection of polynucleotides based on the distance-dependent optical properties of gold nanoparticles. Science 277: 1078–1080. 3. McCormick, R. M., Nelson, R. J., et al. (1997). Microchannel electrophoretic separations of DNA in injection-molded plastic substrates. Anal. Chem. 69(14): 2626–2630. 4. Martynova, L., Locascio, L. E., et al. (1997). Fabrication of plastic microfluid channels by imprinting methods. Anal. Chem. 69: 4783–4789. 5. Duffy, D. C., McDonald, J. C., et al. (1998). Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70: 4974– 4984. 6. Seiler, K., Fan, Z. H., et al. (1994). Electro-osmotic pumping and valve-

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less control of fluid flow within a manifold of capillaries on a glass chip. Anal. Chem. 66: 3485–3491. Kopp, M. U., Mello, A. J. D., et al. (1998). Chemical amplification: continuous-flow PCR on a chip. Science 280: 1046–1048. Wilding, P., Kricka, L. J., et al. (1998). Integrated cell isolation and polymerase chain reaction analysis using silicon microfilter chambers. Anal. Biochem. 257(2): 95–100. Fan, H. A., Mangru, S., et al. (1999). Dynamic DNA hybridization on a chip using paramagnetic beads. Anal. Chem. 71: 4851–4859. Burns, M. A., Johnson, B. N., et al. (1998). An integrated nanoliter DNA analysis device. Science 282(5388): 484–487. Burns, M. A., Mastrangelo, C. H., et al. (1996). Microfabricated structures for integrated DNA analysis. Proc. Natl. Acad. Sci. USA 93(11): 5556–5561. McBride, S. E., Moroney, R. M., et al. (1998). Electrohydrodynamic pumps for high-density microfluidic assays. In: Micro Total Analysis Systems. D. J. Harrison and A. D. Van Den Berg, eds. Banff, Kluwer, pp. 45–48. DeWitt, S. (1999). Microreactors for chemical synthesis. Curr. Opin. Chem. Biol. 3: 350–356. Khandurina, J., Jacobson, S. C., et al. (1999). Microfabricated porous membrane structure for sample concentration and electrophoretic analysis. Anal. Chem. 71(9): 1815–1819. Lucas, F. V., Duncan, A., et al. (1987). A novel whole-blood capillary techniques for measuring the prothrombin time. Am. J. Clin. Pathol. 88(4): 442–446. Gibbons, I., Gorin, M., et al. (1989). Patient-side immunoassay system with a single-use cartridge for measuring analytes in blood. Clin. Chem. 35(9): 1869–1873. Schembri, C. T., Burd, T. L., et al. (1995). Centrifugation and capillary integrated into a multiple analyte whole-blood analyzer. J. Automatic Chem. 17(3): 99–104. Schembri, C. T., Ostoich, et al. (1992). Portable simultaneous multiple analyte whole-blood analyzer for point-of-care testing. Clin. Chem. 38(9): 1665–1670. Deshpande, M., Greiner, K., et al. (1999). Transducers 99, Japan. Ramsey, J. M., Jacobson, S. C., et al. (1995). Microfabricated chemical measurement systems. Nature Med. 1: 1093–1096. Hadd, A. G., Raymond, D. E., et al. (1997). Microchip device for performing enzyme assays. Anal. Chem. 69(17): 3407–3412.

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22. Cohen, A. S., Najarian, D., et al. (1988). Rapid separation of DNA restriction fragments using capillary electrophoresis. J. Chromatogr. 458: 323–333. 23. Cohen, C. B., Chin-Dixon, E., et al. (1999). A microchip-based assay for protein kinase A. Anal. Biochem. 273. 24. Koutny, L. B., Schmalzing, D., et al. (1996). Microchip electrophoretic immunoassay for serum cortisol. Anal. Chem. 68: 18–22. 25. Chiem, N., and Harrison, D. J. (1997). Microchip-based capillary electrophoresis for immunoassays. Anal. Chem. 69: 373–378. 26. Schmalzing, D., Koutny, L. B., et al. (1997). Immunoassay for thyroxine (T4) in serum using capillary electrophoresis and micromachined devices. J. Chromatogr. B. Biomed. Sci. Appl. 697(1–2): 175–180. 27. Li, P., and Harrison, D. J. (1997). Transport, manipulation, and reaction of biological cells on-chip using electrokinetic effects. Anal. Chem. 69: 1564–1568. 28. Fu, A. Y., Spence, S., et al. (1999). A microfabricated fluorescenceactivated cell sorter. Nature Biotechnol. 17(11): 1109–1111. 29. Waters, L. C., Jacobson, S., et al. (1998). Microchip device for cell lysis, multiplex PCR amplification, and electrophoretic sizing. Anal. Chem. 70: 158–162. 30. Waters, L. C., Jacobson, S. C., et al. (1998). Multiple sample PCR amplification and electrophoretic analysis on a microchip. Anal. Chem. 70(24): 5172–5176. 31. Wilding, P., Shoffner, M. A., et al. (1994). PCR in a silicon microstructure. Clin. Chem. 40: 1815–1818. 32. Woolley, A. T., Hadley, D., et al. (1996). Functional integration of PCR amplification and capillary electrophresis in a microfabricated DNA analysis device. Anal. Chem. 68: 4081–4086. 33. Belgrader, P., Benett, W. et al. (1999). PCR detection of bacteria in seven minutes. Science 284(5413): 449–450. 34. Kalinina, O., Lebedeva, I., et al. (1997). Nanoliter scale PCR with TaqMan detection. Nucleic Acids Res. 25: 1999–2004.

3 Short Tandem Repeat Analysis on Microfabricated Electrophoretic Devices Dieter Schmalzing, Aram Adourian, Lance Koutny, Paul Matsudaira, and Daniel Ehrlich Whitehead Institute for Biomedical Research, Cambridge, Massachusetts

3.1 INTRODUCTION Short tandem repeats (STRs) are short repetitive sequences of DNA found throughout the genome [1,2]. They typically consist of seven to 20 iterations of elementary DNA units, each two to seven bases in length. STRs are highly polymorphic, and as a result they are becoming increasingly important for genetic linkage studies, medical applications, parental testing, and determination of personal identity [3]. The forensic community is particularly interested in this system since it provides for a high degree of discrimination between individuals with accurate allele assignments. Forensic STR assays are robust even when degraded DNA or DNA mixtures are used, both common in crime scene DNA evidence. Over the next several years, law enforcement and associated forensic laboratories worldwide will face the challenge of implementing this powerful method for routine use. Such an endeavor involves the establishment of legal procedures and scientific protocols and the creation of large STR databases of convicted felons. 55

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In the United States, the Federal Bureau of Investigation has recently selected a set of 13 standard STR loci for its STR database, called the Combined DNA Indexing System (CODIS) [4]. STR samples are currently analyzed by slab gel or capillary gel electrophoresis after PCR amplification using fluorescent-labeled primers. Several STR loci can be analyzed in a single electrophoretic run when multiplexed by selecting distinct fluorescent labels and varying ranges of fragment sizes for different loci. Allele assignment is achieved by coelectrophoresis of STR standard ladders. Slab gel electrophoresis is inherently slow. A single STR analysis may take several hours. An additional disadvantage is that many operations are difficult to fully automate, particularly gel pouring and sample loading. Capillary electrophoresis potentially allows for complete automation, but still requires several tens of minutes per analysis, thereby limiting throughput [5]. Microfabricated electrophoresis devices, first described by Manz and Harrison [6,7], promise to advance the performance of electrophoretic DNA separations close to theoretical limits. In addition to improved electrophoretic performance quality, such systems are also intrinsically capable of dramatically increasing sample throughput by orders of magnitude, a significant advantage particularly for genotyping applications. These advantages result from the inherent features of microfabricated devices, which allow for the implementation of precisely controlled injection widths, ultrashort separation distances, and very high levels of parallelism. The following sections will describe the general characteristics of this powerful technology and its applications to STR analysis in particular. 3.2

MICROFABRICATION

Most electrophoretic microdevices are made of glass or fused-silica substrates [8,9]. The microfabrication process in such materials is well understood, easily implemented, and extremely versatile. In addition, the chemistries (e.g., for channel wall passivation) originally developed for fused-silica capillaries used in capillary electrophoresis are directly applicable to these substrates.

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Figure 1 depicts the work flow typical for the fabrication of glass and fused-silica devices. First a thin film of metal is deposited onto the substrate, followed by the deposition of a thin film of spin-coated photoresist. The metallic layer serves to improve the adhesion of the photoresist to the substrate wafer during the etching process. Second, the photoresist is patterned by exposure to UV light through a reusable

Figure 1 Work flow of the microfabrication process for electrophoretic microdevices using glass and fused-silica substrates.

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photomask containing the desired channel layout. This is followed by chemical removal of the resist and the metal layer wherever they were exposed to the UV radiation. Third, the resulting pattern is isotropically wet-etched by submerging the wafer in a solution of NH 4 /HF. The isotropy of the etching process produces channel widths of approximately twice the channel depth plus the photomask line width. Longer etching times result in deeper and wider channels. Fourth, resist and metal are chemically removed. Fifth, access holes are drilled into the etched wafer by laser drilling. This is followed by thermal fusion of a second, nonetched fused-silica wafer to the etched substrate wafer, forming a monolithic enclosed channel structure. Finally, vials are attached to the finished microdevice to hold sample and buffer and to allow for electrical contacts to appropriate high-voltage power supplies. Figures 2 and 3 are scanning electron micrographs of the cross section and the top of etched microchannels, illustrating the versatility and the precision of the microfabrication technique. Figure 4 shows an optical photograph of a photomask used to create eight identical eight-

Figure 2 Scanning electron micrograph showing the cross section of a channel wet-etched into a fused-silica substrate. The channel is 28 µm deep and 66 µm wide at the top, yielding a cross-sectional area equivalent to a 44 µm ID cylindrical capillary. (From Ref. 18.)

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Figure 3 Scanning electron micrograph of top view of a channel wet-etched into a fused-silica substrate. The channel dimensions are similar to those in Figure 2.

channel devices simultaneously on a single fused-silica wafer. Resulting individual devices are then cut from the intact wafer with a diamond saw. A photograph of a single-channel microdevice ready for use is shown in Figure 5. Both single-lane and eight-lane device structures have been successfully used for rapid STR analysis. 3.3 MICRODEVICE DESIGN The fundamental channel structure of most electrophoretic microdevices is a simple cross architecture, as illustrated in Figure 6 [10]. The cross consists of a long main separation channel, and three shorter offset side channels which form the injector (‘‘cross channel’’) segment. The separation channel is several centimeters long. The three short side channels are only few millimeters in length, each terminating in either a buffer (B a , B c), sample (S), or waste (W) reservoir. Effective separation channel lengths of only a few centimeters have been found to be sufficient for most STR analysis [11]. All channels have depths of 20–45 µm and widths of 60–100 µm. The micromachined injectors

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Figure 4 Optical photograph of the photolithography mask used for patterning eight identical eight-channel devices simultaneously onto a single circular fused-silica wafer. The individual devices are cut from the bonded wafer prior to use.

have a length of 50–250 µm, determined by the offset between the sample and waste channels. Resulting injection volumes are in the subnanoliter range. Figure 7 shows a schematic of a multiplexed eight-channel STR device for the simultaneous high-speed analysis of eight different STR samples. The structure of the device is an implementation for each separation channel of the simple cross structure described above. The separation distances between injection point and detection point are 4.5– 5.0 cm long. The separation channels are spread out toward the cathodic end to allow for proper arrangement of sample and waste reservoirs. The eight separation channels converge at the anodic end of the device to a density of five lanes per millimeter to minimize the travel distance of the optical laser-induced fluorescence detector. The eight channels share common anodic and cathodic buffer reservoirs (B a , B c). This

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Figure 5 Photograph of a single-channel device (15 cm ⫻ 1.5 cm ⫻ 0.22 cm) made from glass, ready for use. Glass sample and buffer vials (0.5 cm height ⫻ 0.4 cm ID) are affixed onto the microdevice around laser drilled microholes (125 µm in diameter) providing access to the microchannels.

Figure 6 Schematic of a single-channel microdevice. (B a) anodic buffer, (B c) cathodic buffer, (S) sample, and (W) waste. The short channels from the loading channel. The long channel is the separation channel.

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Figure 7 Schematic of an eight-channel microdevice for parallel analysis of eight STR samples. (S n) sample, (W m) waste. (From Ref. 17.)

allows for easy replacement of the polymeric sieving matrix in the entire channel structure. Each separation channel has its own independent sample reservoir (S n), whereas waste reservoirs (W m) are combined whenever possible to economize space and to reduce complexity. Sample and waste reservoirs are connected by loading channels forming offset injector intersections with the separation channels. 3.4

INSTRUMENTATION AND DEVICE OPERATION

A schematic of a single-channel microdevice genotyping apparatus equipped with a four-color laser-induced-fluorescence (LIF) detection system is shown in Figure 8. The system consists of an argon ion laser, a microscope objective to collect the fluorescent emission, a series of dichroic mirrors to separate the fluorescence spectra of up to four different fluorescent dyes, and four photomultiplier tubes (PMTs) to mea-

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Figure 8 Schematic of electrophoretic microdevice apparatus with fourcolor LIF detection system for single-channel analysis.

sure fluorescence signal strengths. The PMTs are interfaced with a computerized data acquisition system. The electrophoretic microdevice is affixed to a temperature-controlled heating block containing a machined aperture to provide optical access to the channel detection zone. A high-voltage power supply is connected via a voltage switching circuit individually addressing the various reservoirs attached to the genotyping device. Once fabricated, the microdevice is prepared for operation by chemically passivating the inner walls of the microchannels to suppress electroosmotic flow and sample adsorption [12]. A polymeric solution (e.g., linear polyacrylamide) is introduced into the entire channel system to provide electrophoretic sieving of the DNA fragments [13]. Buffer is loaded into the buffer and waste reservoirs and a STR sample dissolved in water (2–10 µL) is pipetted into the sample reservoir. A specific voltage sequence is applied to the device and the sample is thereby loaded, injected, and finally separated in the main separation channel. Full analysis of STR samples is completed within minutes. The electrophoretic loading, injection, and separation process is detailed in

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Figure 9 Schematic of the injection segment in load, inject, and run mode, indicating sample (S) and waste (W) reservoirs.

Figure 9. For loading, the sample is electrophoresed from the sample reservoir to the waste reservoir via the loading channel. The voltages are switched from load mode to run mode when a representative aliquot of sample is present in the intersection resulting in the injection of the sample aliquot into the separation channel and the initiation of the electrophoretic separation process. Low positive voltage (‘‘pullback voltage’’) is applied during the run to the sample and the waste channels to prevent leakage of additional sample into the separation channel. Such leakage would cause a significant rise in the electropherogram baseline, complicating interpretation. Identical loading and injection procedures are used in the eight-channel device described previously. The cross injector is a unique feature of electrophoretic microdevices and may offer an important advantage over capillaries whenever electrokinetic injection is employed [14] (pressure injection is not discussed here since it is not applicable to gel based electrophoretic systems). In capillaries, injection of sample is made directly into the separation channel. This limits the time available for injection, since extended injections can lead to large injection volumes, reducing the quality of the separation, while short injections can prevent introduction of slower-moving species. Moreover, electrokinetic injections into capillaries are susceptible to the composition of the sample matrix. For example, high salt content results in reduced voltage drop in the sample vial, yielding less total sample material injected.

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The incorporation of a lithographically defined injection element, the cross injector, in a microdevice offers several potential advantages. First, the arrangement of the loading channel perpendicular to the separation channel allows for loading times as long as necessary to achieve adequate sample loading without affecting the quality of the separation. Second, the fixed injection volume defined by the cross injector likely leads to a significant reduction in the variability and uncertainty of the injection process. 3.5 STR GENOTYPING APPLICATIONS As an example of a two-color multiplexed STR analysis, a typical eight-loci genotyping profile is shown in Figure 10 [15]. These eight STR loci are a subset of the 13 core CODIS loci. The analysis is performed on a single-channel device, similar to the one described above, with an effective separation distance of only 2 cm and using a dualwavelength LIF detection system. The sample is mixed with the allelic standard ladders prior to injection for accurate allele assignment. As is seen, high-quality baseline separation is achieved for all alleles in approximately 2.4 min, allowing for rapid and unambiguous identification of all alleles present in the sample. For example, in Figure 10, the individual is heterozygous for locus D17 (alleles 11 and 13 are present) and homozygous for locus TPOX (only allele 11 is present). The superiority of the microdevice system for the analysis of STRs over more traditional electrophoretic approaches such as slab gels and capillaries is convincingly demonstrated in Figure 11 [16]. Based on analysis time, the microfabricated system outperforms the ABI 373 by a factor of 70 and the capillary system by a factor of ⬎20, with no compromise in data quality. For the microdevice system to become a next-generation technology supplanting existing conventional electrophoresis systems, the ability to increase throughput by multiplexing will also be required. Figure 12 shows a typical genotyping result achieved on a multiplechannel device [17]. The single-color sizing ladder CTTv was used as a test sample and baseline separation was achieved in under 2.5 min in all channels with very high data quality.

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Figure 10 Microdevice electropherograms of the simultaneous two-color analysis of eight STR loci (a) D16S539, D7S820, D13S317, D5S818; and (b) CSF1PO, TPOX, TH01, vWA (CTTv). The allelic standard ladders were mixed with the PCR-amplified sample of an individual before injection. Allele numbers are given above the peaks. Separation conditions: 40-µm-deep channel, 150-µm-long injector, 2-cm separation distance, 50 C, 200 V/cm, 4% (w/v) linear polyacrylamide in denaturing buffer. (From Ref. 15.)

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Figure 11 Comparison of electropherograms of the four loci CTTV STR system of an individual using (a) an ABI 373 slab gel apparatus, (b) a capillary electrophoresis system, and (c) the Whitehead Institute microdevice. The additional peaks of smaller amplitude in (c) arise from the sizing ladder which was mixed with the sample. (From Ref. 16.)

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Figure 12 Multiple-channel genotyping results for the four loci CTTv STR system. Samples were simultaneously cross-injected for 2 min at 200 V/cm and detected in serial with a single-point single-color LIF detector. Separation conditions were as in Figure 10. (From Ref. 17.)

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3.6 OUTLOOK At present, there is no microdevice electrophoresis STR instrument commercially available. However, this exciting technology is under intensive research and development. The first STR analysis instruments may become available to the forensic community within a few years. The acceptance level for this novel technology is likely to be quite high for several reasons. First, the scientific and law enforcement communities are already familiar with the electrophoretic separation processes used in microdevices, which are fundamentally similar to those in conventional slab gels and capillaries. Second, such apparatus should lead to a drastic increase in STR sample throughput, especially significant in the process of establishing convicted felon STR databases. A modestly multiplexed device, such as the eight-channel system described above, would already process 384 multiplexed STR samples in 8 hr, assuming a turnaround time of 10 min when sample handling and reagent replenishment are fully automated. A similar device multiplexed to 96 channels should analyze ⬎4000 STR samples in the same 8-hr period. Third, all the protocols for STR sample preparation and handling already in use for STR analysis by slab gels can be directly transferred to microdevices. Finally, microdevices are inherently amenable to automation and will consequently require significantly less human intervention than traditional methods, resulting in significant savings in labor and cost.

REFERENCES 1. Edwards, A., Civitello, A., Hammond, H.A., and Caskey, C.T. Am. J. Hum. Genet. 49 (1991), 746–756. 2. Beckman, J.S., and Weber, J.L. Genomics 12 (1992), 627–631. 3. Fregeau, C.J., and Fourney, R.M. Biotechniques 15 (1993), 100–119. 4. Parson, W., and Schneider, P.M. Proceedings from the Ninth International Symposium on Human Identification (1998), Promega Corp., Orlando, FL. 5. Lazaruk, K., Walsh, P.S., Oaks, F., Gilbert, D., Rosenblum, B.B., Menchen, S., Scheibler, D., Wenz, H.M., Holt, C., and Wallin, J. Electrophoresis 19 (1998), 86–93. 6. Manz, A., Harrison, D.J., Verpoorte, E.M.J., Fettinger, J.C., Paulus, A., Luedi, H., and Widmer, H.M. J. Chromatogr. 593 (1992), 253–258.

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7. Harrison, D.J., Manz, A., Fan, Z., Luedi, H., and Widmer, H.M., Anal. Chem. 64 (1992), 1926–1932. 8. Garcia Campana, A.M., Baeyens, W.R.G., Aboul-Enein, H.Y., and Zhang, X. J. Microcolumn Sep. 10 (1998), 339–355. 9. Simpson, P.C., Wooley, A.T., Mathies, R.A. Biomed. Microdevices 1 (1998), 7–25. 10. Jacobson, S.C., Hergenroeder, R., Koutny, L.B., Warmack, R.J., and Ramsey, J.M. Anal. Chem. 66 (1994), 1107–1113. 11. Schmalzing, D., Koutny, L., Adourian, A., Belgrader, P., Matsudaira, P., and Ehrlich, D. Proc. Natl. Acad. Sci. USA 94 (1997), 10273–10278. 12. El Rassi, Z., and Nashabeh, W. In: Capillary Electrophoresis Technology, Marcel Dekker, New York (1993), pp. 383–434. 13. Heiger, D.N., Cohen, A.S., and Karger, B.L. J. Chromatogr. 516 (1990) 33–48. 14. Krivacsy, Z., Gelencser, A., Hlavay, J., Kiss, G., and Sarvari, Z. J. Chromatogr. A 834 (1999), 21–44. 15. Schmalzing, D., Koutny, L., Chisholm, D., Adourian, A., Matsudaira, P., and Ehrlich, D. Anal. Biochem. 270 (1999), 148–152. 16. Schmalzing, D., Koutny, L., Adourian, A., Matsudaira, P., and Ehrlich, D. Proceedings from the Eighth International Symposium on Human Identification 1997, Promega Corp. (1998), pp. 112–118. 17. Koutny, L., Schmalzing, D., Adourian, A., Chisholm, D., Matsudaira, P., and Ehrlich, D. Proceedings from the Ninth International Symposium on Human Identification 1998, Promega Corp. 18. Koutny, L., Schmalzing, D., Taylor, T.A., and Fuchs, M. Anal. Chem. 68 (1996), 18–22.

4 Basic Sample Preparation DNA Separation and Purification Rama Ramanujam, Wendy Sacks, and Jie Kang QIAGEN GmbH, Hilden, Germany

4.1 INTRODUCTION Separation and purification of deoxyribonucleic acid (DNA) from other cellular constituents is a prerequisite for most nucleic acid-based drug discovery and clinical diagnostic applications. As with most techniques and tools, a number of options and methods are available for DNA separation and purification, and the optimal method for a particular application must be determined empirically. In recent years, advances in sample preparation techniques for these applications have been achieved by applying the traditional art of chromatography to DNA separation and purification methods. The objective of this chapter is to review selected choices available for the separation and purification of DNA both commercial and otherwise, and discuss the challenges involved in the quest to automate and integrate miniaturized sample analyses. 4.2 CURRENT CHALLENGES Separation and purification of DNA has tended to be practiced as more of an art than a science despite high costs, conventional wisdom, and 71

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suboptimal speed. The challenge is now to keep up with the explosion of genetic information and the advances in information systems. Upstream preparation of high-quality samples must be better, faster, and cheaper to serve the downstream applications spinning genetic information out of nucleic acid–based technologies at ever-accelerating rates. The increase in number of samples and the decrease in analyte size pose a concrete challenge for further innovation and improvement in DNA purification technologies. In drug discovery programs, for example, samples are shrinking in size as they are processed in microtiter plates accommodating from 96 to 384 to 1536 samples and beyond. One approach might well be to find higher-affinity binding supports for nucleic acids than the conventional silica-based gels and ion exchange materials. Another approach might involve improving methods of cell lysis and capture of contaminants such as proteins, lipids, and carbohydrates. The marriage of semiconductor technologies with molecular biology techniques represented by microarray analysis embodies the culmination of the trend toward increasing throughput and diminishing sample sizes. This path, directed toward miniaturized, point-of-care genetic analysis, ultimately demands integrated, automated systems that process and analyze samples from start to finish without human intervention. Hence, the commercial potential and opportunities for developing novel methods and materials for DNA separation and purification continue to be enormous. 4.3

CONVENTIONAL ORGANIC EXTRACTION

The key elements of the DNA purification process are cell disruption and separation of the DNA from other cell components, particularly proteins. Most methods therefore include an anionic detergent such as sodium dodecyl sulfate (SDS) in the initial lysis step to solubilize the cell membrane. The most common method of deproteinizing the cell lysate involves an initial digestion with a proteolytic enzyme such as proteinase K or pronase followed by extraction with phenolchloroform. Finally, the DNA is recovered and concentrated by precipitating with alcohol (usually ethanol or isopropanol) in the presence of

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monovalent cations supplied by adding a salt, such as sodium chloride or sodium acetate. Specifics of the protocol used depend on the starting material for DNA isolation: detergent-mediated lysis is generally sufficient for body fluids and cultured cells whereas specific homogenization and lysis techniques are required for tissue samples, for example. These types of details are provided in subsequent sections. Guidelines for the techniques described above can be found in standard laboratory manuals such as Sambrook et al. [1] and Ausubel et al. [2]. While these types of conventional DNA purification methods can be performed with reagents and supplies that are often readily available in the lab, they do suffer from a number of disadvantages, particularly when large numbers of samples must be handled in parallel. For example, phenol and chloroform are highly toxic and volatile organic solvents that must be handled and disposed of accordingly. Additionally, phenol-chloroform extraction itself involves performing a phase separation in which the aqueous, DNA-containing phase must be carefully pipetted away from the organic phase without disturbing the proteincontaining interface. This procedure, therefore, involves multiple centrifugation steps, after which each sample must be transferred to a new tube. This can be quite tedious when large numbers of samples must be processed. For this reason, it is worth noting that most of the chromatography-based commercial products described below circumvent these safety problems and greatly simplify and streamline the purification process. 4.4 CHROMATOGRAPHIC METHODS The word ‘‘chromatography,’’ meaning ‘‘color writing’’ in Greek, was coined in 1903 by a Russian botanist, Mikhail Semenovich Tswett, for the separation of plant pigments through a chalk column. The art of chromatography relies primarily on the differential solubility and adsorption of compounds to separate molecules between a stationary and a mobile phase. In DNA separation, rapid evolution of modern liquid chromatography has led to the wide selection of stationary materials configured into gels, beads, columns, and membranes. Typically, the gels and beads are packed into columns or utilized in batch procedures for

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chromatographic adsorption and elution of biological materials. Depending on the material, movement of mobile fluid samples through the stationary gels, beads, or membranes may be driven by gravity, centrifugation, or vacuum. Gel- and bead-based chromatographic separation are generally well suited for processing large samples. Since particulate material clogs the stationary purification resin, however, prefiltration or clarification by centrifugation is a necessary first step. The large volumes used generally necessitate a final alcohol precipitation step after chromatography as well. Membrane technology, in contrast, can be incorporated into spin columns or multiwell units designed for vacuum- or centrifugation-driven purification methods well suited to handling large numbers of samples in parallel. Moreover the purification procedures often yield DNA of sufficient concentration for subsequent molecular biology applications without further concentration steps. On the other hand, membrane-based units generally have capacity and structural limitations. 4.5

ANION EXCHANGE CHROMATOGRAPHY

The predominant gel-based chromatographic method for purifying DNA is anion exchange chromatography utilizing diethylaminoethyl (DEAE) as the anion exchanger. The technique makes use of the fact that DNA is a highly negatively charged linear polyanion owing to its phosphate backbone. In liquid anion exchange chromatography, the positively charged ion exchangers bind the negatively charged nucleic acid under low-salt or low-ionic conditions, which is then released through competition using high-salt or high-ionic conditions (Fig. 1). The purified nucleic acid preparation is subsequently desalted for saltsensitive downstream applications. While conventional anion exchangers used for protein purification were based on cellulose, dextran, and agarose with low charge densities, the anion exchangers developed specifically for nucleic acid purification utilize silica beads as the base material which can be charged with a much higher density of DEAE groups. Such high-density anion exchangers offer exceptional separation of all types of DNA (e.g., plasmid, genomic, lambda) because of the wide range of salt concentrations that can be used, effectively purifying away proteins, polysaccharides,

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Figure 1 Simplified interaction between anion exchange material and DNA at different salt concentrations.

metabolites, dyes, oligonucleotides, and ribonucleic acid (RNA) (Fig. 2). The resolving power of some of these anion exchangers is such that it is even possible under certain conditions to separate double- from single-stranded DNA. These types of anion exchange resins are available almost exclusively prepacked in columns as part of ready-to-use commercial kits from a limited number of suppliers (QIAGEN [3], Life Technologies). 4.6 SILICA-BASED METHODS Another chromatographic method for separating and purifying DNA involves using silica (silica particles or slurries, glass powder). In the

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Figure 2 Separation of nucleic acids at neutral pH on QIAGEN anionexchange resin: an example of an anion exchanger designed for nucleic acid purification. (From Ref. 3.)

presence of high concentrations of chaotropic salts, which remove water from hydrated molecules in solution, nucleic acids adsorb to silica, while molecules such as polysaccharides and proteins do not adsorb and are removed. This method has the advantage that DNA is eluted under low- or no-salt conditions, which is ideal for salt-sensitive downstream applications. One disadvantage of this technique, however, is that silica fines may contaminate the final preparation, which can inhibit certain enzymatic reactions. A number of suppliers offer columns with integrated silica membranes, which circumvent this problem. These include QIAGEN and Promega, among numerous others. Most products are designed for isolating specific types of DNA (e.g., plasmid or genomic) from particular sample sources (e.g., bacteria, clinical samples, or plants) and some are designed for specific downstream applications, such as PCR.

4.7

PLASMID DNA

Plasmid DNA plays an important role in molecular biology and genetic engineering research. A myriad of plasmid purification options have been developed ranging from simple lysis to complex chromatographic techniques. The most common method for partially purifying plasmid

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DNA is based on the alkaline lysis procedure as described by Birnboim and Doly [4]. In brief, the alkaline lysis protocol utilizes SDS to lyse bacterial cells and NaOH to denature DNA and protein. On neutralization with acidic potassium acetate, chromosomal DNA, proteins, and cellular debris become entrapped in precipitated salt-SDS complexes. Plasmid DNA, on the other hand, remains in solution since it is relatively small and covalently closed and therefore renatures readily. Centrifugation removes the precipitated contaminants. The plasmid DNA in the supernatant can either be collected and concentrated by alcohol precipitation or used as the starting material for further purification methods such as those described below. Historically, when larger quantities of plasmid DNA (hundreds of micrograms and up) of the highest purity were required, the most common purification method used was cesium chloride-ethidium bromide gradient centrifugation [1,2]. However, this method suffers from a number of disadvantages including technical difficulty, use of a carcinogen (ethidium bromide), and requirement for specialized equipment (an ultracentrifuge and appropriate accessories), as well as being quite time-consuming (requiring anywhere from 1 to 3 days). Commercial kits utilizing anion exchange columns have therefore become the standard for larger-scale plasmid purification since they circumvent all of these limitations while yielding DNA of at least comparable quality [5]. For purification of smaller quantities of plasmid DNA from larger numbers of samples, the state of the art in terms of commercial products are those employing silica membrane technology because of their speed, handling convenience, availability in multiwell formats, and cost-efficiency. Alternatively, phenol-chloroform extraction also continues to be used for smaller plasmid preparations. 4.8 GENOMIC DNA TEMPLATES FOR PCR The explosion of PCR [6,7] into molecular biology, molecular medicine, routine diagnostics, forensics, epidemiology, ecology, and breeding, to name only a few applications, has led to the need for DNA purification both before and after amplification. However, the diversity

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of the sources of PCR templates for these applications—whole blood, buccal swabs, mouse tail, soil samples, plant material, animal tissues, cultured cells, yeast, and bacteria, for example—preclude a single protocol being effective for all material. Instead, protocols must be optimized for the specific application, particularly with respect to sample lysis and homogenization. A few major sample types are considered below; a number of protocols for other sample types can be found in general PCR reference works [e.g., 8,9] as well as general laboratory manuals [e.g., 1,2]. Though PCR amplification is permissive to ‘‘quick and dirty’’ sample preparations, in most situations, good-quality DNA is required for successful and reproducible results. In addition to components of specific sample types that inhibit PCR (e.g., heme; see specific sections below), many reagents used routinely in traditional DNA purification methods can inhibit the reaction, even at low concentrations (Table 1). Proteinase K, commonly used in genomic DNA purification protocols, Table 1 Impurities Showing Inhibitory Effects on PCR Substance inhibitory concentration SDS ⬎0.005% (w/v) Phenol ⬎0.2% (v/v) Ethanol ⬎1% (v/v) Isopropanol ⬎1% (v/v) Sodium acetate 3 5 mM Sodium chloride 3 25 mM EDTA 3 0.5 mM Hemoglobin 3 1 mg/mL Heparin 3 0.15 IU/mL Urea ⬎20 mM Reverse transcription reaction mixture 3 15% Concentrations of impurities showing inhibitory effects on PCR amplification were determined in eight different PCR systems for all substances except urea and heparin, which were tested in four different systems. Source: Ref. 10.

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must be thoroughly inactivated or removed, or it will digest the thermostable DNA polymerase used for amplification. In spite of all of the disadvantages described earlier, organic extraction remains a common means of purifying genomic DNA. It should be noted that a limited number of suppliers offer commercial products that provide a number of safety and handling advantages as well as removing or avoiding the use of the PCR inhibitors mentioned above. 4.9 BLOOD AND BODY FLUID SAMPLES Blood samples provide an invaluable source of nucleic acids for genetic characterization and are therefore a very common source of template DNA for PCR. However, blood samples are specifically associated with several known PCR inhibitors. The heme component of hemoglobin is known to inhibit the reaction [11] as are two common anticoagulants used to treat blood samples: heparin and EDTA (Table 1). Nevertheless, there are a number of published purification protocols that yield genomic DNA from blood suitable for PCR amplification. One of the most common is the ‘‘salting out’’ method [12]. Other simple nonorganic procedures involve lysis, centrifugation of nuclei, and treatment with proteinase K followed by inactivation of the enzyme [13,14]. In one such protocol, one volume of whole blood is mixed with 10 volumes of TE (10 mM Trisu`Cl, ly¨mM EDTA, pH 7.5 or 8.0) in a microcentrifuge tube and centrifuged for 10 sec at 13,000g. The pellet is then resuspended in the same volume of TE, mixed, and centrifuged as before. This procedure is repeated two more times before resuspending the pellet in two volumes (relative to the starting volume) of PCR buffer lacking gelatin or bovine serum albumin but containing 1% Laureth 12 or 0.5% Tween 20, and 100 µg/ mL fresh proteinase K. The mixture is incubated at 56°C for 45 min followed by a 10-min incubation at 95°C to inactivate the protease. Ten microliters can be used in PCR [14]. As noted above, ready-to-use and optimized products employing spin columns, such as QIAamp Kits from QIAGEN, can be used to purify genomic and viral DNA from fresh, frozen, or dried whole blood

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(collected in citrate, EDTA, heparin, or other common anticoagulants), as well as other body fluids, cells, and tissues. The DNA is suitable for amplification and blotting applications ranging from HLA typing, paternity testing, bacterial diagnostics, human and animal testing, viral diagnostics, and forensic analysis to cancer research and transgene screening.

4.10 MAMMALIAN TISSUE SAMPLES Genomic DNA can generally be isolated from small amounts of softtissue samples as described above for body-fluid samples. Hard-tissue samples, on the other hand, often must be thoroughly ground (for example, in liquid nitrogen) and/or extensively deproteinized in order to extract good-quality genomic DNA. Depending on the material, most standard protocols include a proteinase K digestion ranging from 1 to 24 hr as well as extensive organic extraction followed by alcohol precipitation. Both standard laboratory manuals [e.g., 1] and general PCR guides [e.g., 8,9] provide a variety of such protocols for different applications. Once again, far more convenient, safer, and reliable alternatives utilizing spin columns or multiwell purification units are offered by the commercial suppliers mentioned above.

4.11 PLANT MATERIAL Plants present special challenges to researchers attempting to extract nucleic acids. The diversity both among the species themselves (from weeds to food crops to trees) and among the different tissue types (from seeds to leaves to flower parts) makes it difficult to develop methods that function universally. In addition, plants often accumulate large quantities of materials that complicate DNA isolation and interfere with PCR and other enzymatic reactions, such as polysaccharides, nucleases, and a variety of metabolites, to name but a few. Many common plants contain polyphenolics such as terpenoids and tannins which bind to nucleic acids, reducing DNA quality [15]. Physically resistant constituents such as the cell wall, moreover, mean that specific steps must often be taken to disrupt, lyse, and filter plant material.

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Plant tissue is most commonly disrupted by freezing in liquid nitrogen and grinding, for example, in a mortar and pestle, although a common alternative is lyophilization prior to grinding. Because it is generally effective for a wide variety of species and tissue types, a classical method of preparing plant DNA involves extraction with the nonionic detergent cetyltrimethylammonium bromide (CTAB) in the presence of high concentrations of NaCl. Inclusion of a chloroform extraction step under these conditions specifically removes polysaccharides. DNA is recovered finally by alcohol precipitation [16]. A frequently cited miniprep method involves extraction in SDS and potassium acetate precipitation, followed by multiple rounds of alcohol precipitation [17]. Alternatively, standard organic extraction procedures are also commonly used as described above. Polyphenolic compounds can be removed by including 1% polyvinylpyrrolidone (PVP) during homogenization, yielding DNA suitable for restriction digests, Southern analysis, PCR amplification, and library construction [16,18]. While the methods described above are quite versatile, the number of manipulations make them impractical for high-throughput analyses. Many protocols have subsequently been published in which the entire procedure can be carried out in a single tube or well without any centrifugation steps. Two examples of these utilize Tris-EDTA-lauroyl sarkosyl extraction buffers containing either proteinase K [19] or polyvinylpolypyrrolidone (PVPP) [20] and yield DNA suitable for PCR and RAPD analysis. Among the very limited number of commercial products for plant DNA purification on the market, only the DNeasy Plant System from QIAGEN employs spin column or multiwell plate technology. 4.12 PCR FRAGMENTS Depending on the subsequent application, amplified PCR fragments must be purified away from reaction components such as primers, nucleotides, enzymes, mineral oil, and salts. Any number of methods may be used for this purpose, including simple alcohol precipitation, organic extraction, or purification using silica-based methods [21]. Agarose or polyacrylamide gels are frequently used to separate the specific PCR product from non-specific or undesired products,

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which then requires that the gel material, ethidium bromide, dyes, and sometimes detergent be subsequently removed. One common method is to use chaotropic salt to solubilize the agarose and bind the DNA to silica material [22]. Other methods include electroelution and the use of low-melt agarose [2]. In addition, a large number of companies offer reagents and kits that efficiently remove all of these contaminants using resins, silica slurries, or silica membrane technology [23]. 4.13 AUTOMATION When samples shrink in size and increase in numbers, it is imperative that repetitive and sequential processing steps be handled rapidly and accurately. This requirement highlights the importance of advances in robotics systems and computer networks. Scientists look to automation for simplicity, throughput, and ease of use. They ultimately expect a ‘‘black box’’ that can purify samples unattended, from the initial reagent introduction to collection of the final preparation for downstream applications.

Figure 3 The BioRobot 9600 is a laboratory workstation for automated molecular biology applications.

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Rising to the demand, instrument manufacturers such as QIAGEN, Tecan, and Beckman have developed workstations that automate nucleic acid purification procedures. These robotic systems include various combinations of software programming, reagent delivery, micro-/ nanoliter liquid dispensing, carousel-fed plate movement, agitation units, cooling and heating platforms, vacuum stations, and robotic pickand-drop arms. While not yet the all-encompassing black boxes described above, these robotic systems offer the benefit of performing repetitive, error-prone tasks with greater speed, accuracy, and convenience. Among the work stations available, those from QIAGEN probably provide the broadest range of automated nucleic acid purification and molecular biology applications (Fig. 3). 4.14 DNA QUALITY FOR CHIPS AND ARRAYS Silicon or glass slides bearing anywhere from hundreds to hundreds of thousands of immobilized snippets of DNA will become the microchips of the 21st century [24]. Such chips carrying arrays of cDNAs, PCR fragments, or oligonucleotides will serve as probes for detecting complementary nucleic acid sequences within cDNA or amplified genomic DNA samples. These microchips should ultimately enable researchers to analyze large numbers of genes for diagnostic and therapeutic applications. The purity of the nucleic acids used for microarray analysis is thought to be especially critical to the success of the technique, since it generally demands quantitative hybridization with minuscule amounts of immobilized DNAs, possibly of limited length (in the case of oligonucleotides), within minute areas. The demanding nature of these conditions means that the effects of contaminants of any sort both on hybridization and detection (e.g., autofluorescence of impurities) will be magnified. Nucleic acid isolation or cleanup is therefore generally necessary during both phases of the process: array construction and sample preparation for hybridization. Chips are most frequently spotted with PCR products, whether the original templates are cDNAs, expressed sequence tags, or genomic sequences. Effective automated high-throughput methods for cleanup of PCR products are clearly a requirement for this application. Sample prepara-

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tion for hybridization usually involves several steps, most of which also require nucleic acid purification. For expression analysis applications, highly pure total RNA or poly A⫹ mRNA must be isolated and subsequently reverse-transcribed, possibly amplified, and labeled. Genotyping applications currently focus on the presence or absence of specific sequences in a particular sample rather than whole-genome analysis and therefore involve amplification of the relevant sequences from genomic DNA templates. Reliable, high-throughput methods are therefore required for genomic DNA and RNA/poly A⫹ mRNA isolation, as well as PCR/labeling reaction cleanup. For all of these microarray applications, commercial products employing silica-gel-membrane technology (particularly those from QIAGEN) are recommend by chip suppliers because of their efficacy and reliability, as well as the possibility of automating these procedures. ACKNOWLEDGMENTS We would like to thank Ulrich Schriek and Joanne Crowe for critical reading of the manuscript. REFERENCES 1. Sambrook, J., E. G. Fritsch, and T. Maniatis. 1989. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. 2. Ausubel, F. M., et al. (eds.). 1997. Current Protocols in Molecular Biology. John Wiley and Sons, New York. 3. QIAGEN Inc. The Art of Plasmid Preparation. Santa Clarita, CA, p. 26. 4. Birnboim, H. C., and J. Doly. 1979. Nucl. Acids Res. 7: 1513–1523. 5. Schleef, M., and P. Heimann. 1993. BioTechniques. 14: 544. 6. Saiki, R. K., S. Scharf, F. Faloona, K. B. Mullis, G. T. Horn, H. A. Erlich, and N. Arnheim. 1985. Science 230: 1350–1354. 7. Mullis, K. B., and F. A. Faloona. 1987. Methods Enzymol. 155: 335– 350. 8. Innis, M. A., D. H. Gelfand, J. J. Sninsky, and T. J. White (eds.). 1990. PCR Protocols. A Guide to Methods and Applications. Academic Press, San Diego.

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9. McPherson, M. J., P. Quirke, and G. R. Taylor (eds.). 1991. PCR: A Practical Approach. IRL Press, New York. 10. QIAGEN Inc. Critical Factors for Successful PCR. QIAGEN, Santa Clarita, CA, p. 25. 11. Walsh, P. S., D. A. Metzger, and R. Higuchi. 1991. BioTechniques 10: 506–513. 12. Miller, S. A., D. D. Dykes, and H. F. Polesky. 1988. Nucleic Acids Res. 16: 1215. 13. Grimberg, J., S. Nawoschik., L. Belluscio., A Turck., and A. Eisenberg. 1989. Nucleic Acids Res. 17:y¨8390. 14. Kawasaki, E. S. 1990. In: Innis, op. cit. PCR Protocols, pp. 146–152. 15. John, M. E. 1992. Nucleic Acids Res. 20: 2381. 16. Ausubel. Curr. Protocols. 2.3.1–2.3.7. 17. Dellaporta, S. L., J. Wood., and J. B. Hicks. 1983. Plant Mol. Biol. Rep. 1: 19–21. 18. Pich, U., and I. Schubert. 1993. Nucleic Acids Res. 21: 3328. 19. Guidet, F. 1994. Nucleic Acids Res. 22: 1772–1773. 20. Steiner, J. J., C. J. Poklemba, R. G. Fjellstrom, and L. F. Elliott. 1995. Nucleic Acids Res. 23: 2569–2571. 21. Ausubel. Curr. Protocols. 2.1.1–2.1.9. 22. Coleman, A., M. J. Byers., S. B. Primrose, and A. Lyons. 1978. Eur. J. Biochem. 91: 303–310. 23. Mack, A. 1996. The Scientist. 10: 17–18. 24. Leadon, S. A., and P. A. Cerutti. 1982. Anal. Biochem. 120: 282–288.

5 Multiplexed Integrated Online Sample Preparation for DNA Sequencing and Genetic Typing Edward S. Yeung Iowa State University, Ames, Iowa

5.1 INTRODUCTION The Human Genome Project is an initiative to sequence the entire human genome, which consists of 3 ⫻ 109 bp of nucleic acids. The magnitude of the problem is immense. Recent developments have shown that production centers can be set up to scale up current technology substantially in terms of speed and throughput [1]. Still, one needs a major departure from current technology to go the next step in sequencing the human genome economically and in a timely manner. This will become even more critical as we start to use the information clinically, when the process will have to be repeated for every patient, or in biology, when interspecies and intraspecies variations provide some of the most interesting insights. It is important to note that an integrated approach is critical to success. What works best when small numbers of samples with few time constraints are sequenced may not be transferable to a large-scale project. It may therefore be necessary to reoptimize the performance of each of the critical technologies to achieve a workable compromise for large-scale applications. One does not necessarily strive for the best 87

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detection limits, the most efficient separations, the highest degree of flexibility, or the most sophisticated hardware and software. There is little to be gained if reducing the cost of some of the steps leads to incompatibility with high-speed, high-throughput operation, and ultimately higher costs for the other steps. We need, however, a set of technologies that can function in concert through the entire operation to achieve the final goal of cost-effective, high-speed, high-throughput DNA sequencing. It is also obvious that irrespective of whichever basic technology is eventually employed to sequence the entire human genome, there are substantial gains to be made if a high degree of multiplexing of parallel runs can be implemented. Such multiplexing should not involve expensive instrumentation and should not require additional personnel, or else the main objective of cost reduction will not be satisfied even though the total time for sequencing is reduced. Of all the alternative technologies being developed for DNA sequencing, electrophoresis in multiplexed capillary tubes or microchannels appears to be most ready for real applications in high-speed, high-throughput production environments. The projected cost in the separation (electrophoresis) and the identification (base-calling) tasks of DNA sequencing is expected to be substantially lower than current sequencers. This is a direct result of automation (reduced labor costs) and highly multiplexed operation (reduced instrument cost per called base). So far, a third feature of capillary or microchannel electrophoresis has not been exploited for cost reduction in DNA sequencing. This has to do with the small volumes and small amounts of samples that are needed for each electrophoretic run. Typically, only 10–20 nL of a given sample is injected into the capillary tube or microchannel at concentrations and compositions identical to those loaded into wells in the commercial sequencing instruments. To inject the samples, however, much larger volumes (at least 1 µL) are needed for handling. Furthermore, sample preparation by robotic work stations also requires solution volumes in the microliter range. One must therefore miniaturize the sample preparation (front-end) steps before capillary electrophoresis and to interface these directly to the capillary array in order to take

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advantage of the much lower sample requirement. Substantially lower costs will then be achieved through a true reduction in the amounts (but not necessarily the concentrations) of enzymes and labels used, the amount of bacterial cells needed, and a reduction in labor and instrument costs through automation and multiplexing. There is also the potential for further reductions in costs even in the many steps before cell cultures. It is therefore important to develop front-end work, from harvesting plasmid DNA from cells to producing fluorescence-labeled Sanger fragments, so that potentially sample preparation can be reduced in volume by 100-fold, increased in speed by 10-fold, and multiplexed up to 1000 different samples at a time. These new technologies will be able to interface directly with any microchannel or capillary array instrumentation, so that immediate benefits to the Human Genome Project are foreseen. It is the recent advances in capillary array electrophoresis in our and other laboratories that is the main incentive for such new technologies, since traditional sample preparation was not designed to benefit from the potential speed and cost advantages offered by microchannels or capillary electrophoresis. Conversely, there is no reason to miniaturize and multiplex sample preparation if instrumentation is not available to separate and detect the Sanger fragments afterward. At present, highly multiplexed CE imposes a great demand on the throughput of sample preparation for DNA sequencing. Linear amplification of sequencing (cycle sequencing), which is the Sanger dideoxy termination chemistry modified by the concept of polymerase chain reaction (PCR), has gained popularity due to the lower DNA template requirement, ease of automation, and effective denaturation of double-stranded DNA [2,3]. The marriage of automated cycle sequencing with highly multiplexed capillary array electrophoresis has the potential to further increase the throughput and reduce the cost in large-scale DNA sequencing projects. The small sample requirement of CE, being as low as ⬃5 nL, provides a great opportunity to reduce the amount of DNA template and reagents, leading to a substantial reduction of the cost per base unit. Miniaturization of cycle sequencing in a glass capillary has been

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demonstrated [4,5]. PCR reaction in a capillary format has recently been achieved in only ⬃1.5 µL of total volume [6]. Moving the cyclesequencing reaction into a capillary has the additional advantage of increasing the reaction speed due to the small heat capacity of a capillary versus a heating block or a water bath. A capillary reactor is also compatible with highly multiplexed electrophoresis in a parallel capillary array. A microchip providing integrated operation from Sanger reaction to sequencing separation is a promising approach, but such has not yet been demonstrated. Numerous endeavors have been made on developing robotic work stations to perform the sequencing reaction, purification, preconcentration, denaturing, and sample loading in slab gel electrophoresis (SGE) [7–9]. Although robotization has shown advantages in repetitive operation with high precision, the adaptation to highly multiplexed capillary array separation and detection suffers from many incompatibilities in terms of the total reaction volume, purification by centrifugation, lyophilization, and sample injection after reconstitution and denaturation. Many DNA purification methods such as ultrafiltration, spin-column gel filtration, acetate-ethanol precipitation, and phenol-chloroform extraction require a centrifuge to remove fluorescently or radioactively labeled primers or terminators in cycle-sequencing strategy. Interfacing the centrifuge into the robotic work station complicates the whole sequencing protocol. A vacuum chamber under the microtiter plate has been used to replace the centrifuge in a robotic work station [10]. Solid-phase magnetic beads have also been utilized to bind the cyclesequencing products [11,12]. Almost all these modifications still depend on the complicated combination of multiple movements of a robotic arm and precise liquid dispensing on the robotic platform. The inability to miniaturize the robotic work station restricts further miniaturization and multiplexing of the sequencing reactor to couple to CE arrays. An alternative concept to purify the sequencing reaction products is online chromatographic separation. Size exclusion chromatography (SEC) [13] has the unique features of high recovery, desalting ability, favorable elution order for subsequent injection and cleanup, and suitability for pressure flow instead of centrifugation. The online coupling of SEC with CE should

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also allow multiplexed injection of Sanger reaction products into a CE array. 5.2 ONLINE SANGER REACTION AND DNA SEQUENCING The ultimate goal in the Human Genome Project is to design an instrument which can perform DNA sequencing in an online format, starting from cells, through sequencing reaction, online purification, online denaturation, online injection, capillary electrophoretic separation, and ending with called bases. Such a system should also be compatible with a high degree of multiplexing. The important first step is to show that an integrated online system is feasible. We have studied the critical parameters in each of the many steps from template to called bases [14]. An optimized combination is demonstrated, albeit in a manual mode of operation and with bulky switching valves and connectors. Eventual automation and adaptation to a multiple-capillary array system should allow high-speed, high-throughput DNA sequencing from templates to called bases in one step. Figure 1 is a schematic of the entire instrumental setup. A dyelabeled terminator cycle-sequencing reaction is performed in a fusedsilica capillary MR. Subsequently, the sequencing ladder is directly injected into a size exclusion chromatographic column PC operated at ⬃95°C for purification. Online injection to a capillary for electrophoresis SC is accomplished at a junction set K at ⬃70°C. High temperature at the purification column and injection junction prevents the renaturation of DNA fragments during online transfer without affecting the separation. The high solubility of DNA in and the relatively low ionic strength of 1⫻ TE buffer permit both effective purification and electrokinetic injection of the DNA sample. The system is compatible with highly efficient separations by replaceable poly(ethylene oxide) (PEO) polymer solution in uncoated capillary tubes. One of the potential problems is that a large surface-to-volume ratio can inhibit DNA amplification due to adsorption. To recover the activity of these polymerases, bovine serum albumin (BSA) has been found effective in inhibiting adsorption [15]. Preferential adsorption of

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BSA onto the capillary wall protects the polymerase from denaturation. To perform cycle sequencing in a glass capillary requires a vapor-tight system. If the capillary is used with the ends open, a serious problem observed is the segmentation of the solution plug inside the capillary due to nonuniform heating. We found that pressurization of the capillary microreactor under 20–30 psi effectively suppressed segmentation of the solution plug. Sequencing reactions were performed inside a hot-air thermal cycler. Modifications were made to fit the protocol to a capillary format since the original protocol was developed for the ABI model 9600 or 2400 instrument. BSA was added to prevent denaturation of the enzymes on the surface of capillary tubes [16]. The microreactor MR was a 60-cm-long, 250-µm-ID, 360-µm-OD fused-silica capillary. The cycle-sequencing reaction mixture was loaded into the capillary microreactor at port B3 by vacuum produced by a 100-µL glass syringe S4 at port A6. Before temperature programming began, the microreactor was pressurized under 20–30 psi He by selecting port A5 and B4. After the sequencing reaction was complete, chromatographic injection via transfer line T1 was accomplished by pressurizing the products at 30 psi for 1 min at port A5. A µLC-500 pump (ISCO) was used to deliver an elusion buffer to the purification column. The purification column was prepared from 31-cm-long, 0.04-in.-ID, 1/16-in.-OD PEEK tubing. The packing material was superfine G-25-50 Sephadex

Figure 1 Schematic of instrumental setup for integrated online cycle-sequencing SEC-CE system. A, B, and D: six-position selection valves; C: dualposition switching valve; E: needle valve; S1: 1 ⫻ TE buffer; S2: methanol; S3: deionized water; S4: dry and clean syringe; S5: 1 M NaOH; W1–5: waste outlets; AF: in-line filter unit; T: PEEK tee; MR: microreactor capillary; T1 and T2: transfer capillaries; PC: purification column; TC: hot-air thermal cycler; P1: 5 mg/mL BSA solution; P2: deionized water; P3: 1 ⫻ TBE buffer; K: PEEK cross; HB: hot-water bath; G: grounded stainless-steel tubing; SC: separation capillary; L1 and L2: lenses; MO1 and MO2: microscope objectives; IF: 500-nm long-pass filter; BS: beam splitter; LPa: 2 ⫻ 530-nm longpass filters; LPb: 2 ⫻ 610-nm long-pass filters; LPc: 610-nm long-pass filter; PMT-a, b, c: photomultiplier tubes; and A/D: DT2802 A/D interface. (From Ref. 14.)

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particles. The spherical particles have 20–70 µm of swollen bead diameter with useful fractionation range from 1000 to 5000 MW for globular molecules, which should retain excess dye-labeled dideoxyribonucleotides but exclude the other, longer DNA fragments. A PEEK cross K connected the transfer line T2 with a polymerfilled separation capillary (SC) in the opposite direction, with the other two arms forming a flow channel. Syringes P1-3 contained 5 mg/mL BSA aqueous solution, deionized water, and 1⫻ TBE buffer, respectively. The 1⫻ TBE buffer syringe served as the electrophoresis buffer reservoir. The whole junction was immersed in a grounded hot water bath held at 65–75°C, but the outlet of the stainless-steel tubing was kept away from the water bath to avoid forming a short circuit. Before electrokinetic injection, the cross region was rinsed with at least 1 mL of BSA solution, followed by the same amount of deionized water. When PMT-c indicated that the first peak from the size exclusion column reached the cross junction, injection was initiated at 6 kV for 30 sec at the leading edge of this peak. The transverse flow stream was completely stopped by needle valve E during injection. After injection, the effluent from the purification column was directed to the waste line W5 so that the back pressure from the transfer line was released immediately. The needle valve E was reopened and a total of 0.5–1 mL 1 ⫻ TBE buffer solution was allowed to flush slowly through the cross junction before electrophoretic separation. Figure 2a shows the electropherogram of M13mp18 DNA template cycle sequenced by Thermo Sequenase in a pressurized fusedsilica capillary [14]. After a standard cycle-sequencing reaction followed by an optimized cleaning process, a sequencing reaction mixture without DNA template was re-loaded into the capillary microreactor. The reaction products, purified by either a spin-column or homemade Sephadex column, were then injected into a separation capillary to check for cross-contamination. A typical electropherogram of these purified samples is shown in Figure 2b. In this example, interference to the sequencing run, albeit minor, occurs within a few minutes after the first eluted peak. In the actual online system discussed below, we did not observe even the small interference peaks as a result of the longer purification column and the smaller residual amounts of dye terminators. This suggests that the cleaning procedure does effectively remove

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Figure 2 (a) Electrophoretic separation of DNA fragments after cycle-sequencing reaction for M13mp18/U ss-DNA templates amplified by Thermo Sequenase in a fused-silica capillary microreactor. (b) Electropherogram for a blank cycle-sequencing reaction (without DNA template) under the same conditions as (a) and in the same capillary after extensive flushing. Both electropherograms are plotted on the same scale. (From Ref. 14.)

all interference from the previous reaction. This shows that it is possible to reuse the microreactor, which is essential for multiplexed highthroughput operation envisioned for the Human Genome Project. It is interesting that resolving the nested fragment set from the labeled ddNTPs in a low ionic strength solution cannot be achieved. By replacing the 1/100 ⫻ TBE buffer with a 1 ⫻ TE buffer to increase the ionic strength, it is clear that one can obtain baseline-resolved separation and improved peak symmetry for both peaks. The potential drawback of using 1 ⫻ TE buffer is the loss of the benefit of preconcentration by stacking during injection of the Sanger products later on.

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Recycling a purification column to lower operating cost is not often done in a molecular biology laboratory. However, this will be an important issue in a multiplexed, high-throughput operation. In this design, the purification column was placed in a hot-air thermal cycler, which promotes self-cleaning. A typical terminator-labeled cyclesequencing reaction is done in ⬃2.5 hr, and microreactor regeneration, sample loading, and filling of the separation matrix require ⬃0.5 hr. The 3 hr, 10-µL/min flow gives ⬃1800 µL of eluent, corresponding to six packed-column volumes. So, the column can be flushed during the sequencing reaction. We showed that carryover in the purification column is not a problem. By examining the peak widths after the purification column (at PMT-c), one can conclude that the sample plug from the microreactor was diluted by at least a factor of 10 during purification based on the peak elusion volume of ⬃50 µL. Also, typical DNA sequencing samples are obtained in dry form and are redissolved in a 4-µL formamideEDTA solution before denaturation and injection. If instead our product mixture was used directly for injection (without preconcentration), another dilution factor of 5 exists because the standard cycle-sequencing reactions produce a total volume of 20 µL. So, the net dilution factor is ⬃50. Therefore, a concern is how to inject a detectable quantity of diluted DNA sample into CE. Surprisingly, we found that the injection of DNA samples diluted by the low-ionic-strength solutions can tolerate up to 100-fold dilution without noticeable changes in S/N ratios and separation resolution when the injection voltage or injection time was increased. The effect of the low ionic strength buffer on injection is presumably attributed to electric field amplification [17–19]. However, the samples diluted with 1 ⫻ TE buffer (which has higher ionic strength) only tolerated up to 20-fold dilution, producing a decrease in S/N ratio by a factor of 5. Fortunately, the sensitivity of the detection system is more than enough to allow such a decrease in S/N (vide infra). Chemicals, salts, and heat are the common agents for denaturing DNA. In standard DNA sequencing protocols, the purified dry sequencing products are heated in a denaturing solution at 95°C for a few minutes, and then loaded into the slab gel. However, online denaturation may not be necessary because the purification column is operated at

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very high temperature and because of relatively fast sample transfer. The use of high temperature to denature DNA is very common in PCR. This led to the implementation of ‘‘hot injection’’ to guarantee that the fragments remain denatured. The whole cross junction was placed in a heated water bath. The purified sample was sent from the transfer capillary T2 into the cross region without the addition of any formamide-EDTA solution. It was found that the optimal temperature for denaturation of pGEM or M13mp18 DNA is 65–75°C. If the temperature is too low, the DNA sample does not completely denature, but if the temperature is too high, bubbles form easily at the tip of the capillary. The dilute DNA sample (compared to those loaded into slab gels) also minimizes problems associated with renaturation. Electropherograms from online cycle sequencing, purification, injection, and separation of a M13mp18 sample were recorded by using one-wavelength excitation and dual-wavelength detection [20]. Data in both channels show very high S/N ratios, without any interference from dye-labeled ddNTPs. Resolution factors at ⬎0.5 can still be seen toward the end of the separation. By using in-house software [21], we were able to call the sequence with an accuracy of 98% through 370 bp. For three consecutive runs of M13mp18 ss-DNA template in the complete system, the retention time, signal intensity, and resolution of separation were comparable to the variations in the offline separations. pGEM ds-DNA template was tested right after an online run for M13mp18 template. The correct sequence of pGEM DNA can also be called. This indicates that the carryover from run to run in the online system is negligible. 5.3 ONLINE PCR ANALYSIS AND GENOTYPING We have also demonstrated an integrated online system for PCR and capillary gel electrophoresis (CGE) for DNA typing and disease diagnosis. There is no need to reiterate the development of CGE as a powerful analytical tool in post-PCR analysis. These include high-speed, high-resolution restriction fragments analysis [7,22–26], rapid and precise DNA typing and sizing [27–33], single-base mutation analysis [34–39], and the analysis of disease causing genes [40–43]. In particular, capillary array electrophoresis along with other microfabricated

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devices [44–46] are promising methods for the purpose of achieving high-throughput DNA analysis. The ultimate goal is to devise an automatic system which can integrate all these steps, from a real biological sample to final readable results. Two applications have been investigated: the four short tandem repeat (STR) loci vWA, THO1, TPOX, and CSF1PO (CTTv) for DNA typing, and DNA probe for human immunodeficiency virus (HIV-1) diagnosis. The CTTv are important loci in forensic and genetic linkage analysis. The PCR technique can detect the presence of the virus before any antibody response is detectable in the infected person. We have modified the above system to allow one-step ss- and dsDNA fragment analysis after PCR [47]. Modifications were made both in the amounts of PCR reagents and the temperature and time protocol to fit a hot-air thermal cycler. Some research done in the investigation of PCR in microfabricated chips showed that native silicon was an inhibitor of PCR. Amplification in an untreated chip had a high failure rate [48,49]. Complicated surface treatments are necessary such as silanization followed by coating of a selected protein or polymer and the deposition of a nitride or oxide layer onto the silicon surface. We found that high concentrations of BSA in the PCR mixture can prevent surface inhibition. No surface modification was needed for fused-silica capillary as a reactor if BSA was present in PCR mixture. Multiplexed PCR in a fused-silica capillary, online injection, DNA denaturation, and calibration based on a standard ladder have been successfully combined. Also, online liquid flow management, DNA separation, and detection have been completely integrated. Simultaneous amplification of more than one DNA region of interest in one reaction mixture reduces labor, time, cost, and crosscontamination, since sample handling is minimal. For multiplexed amplification, there is competition among the four pairs of primer. There are differences in the optimum conditions for the individual loci. Therefore, a multiplex system has much narrower tolerance limits than single PCR systems. Many factors may affect the amplification results, such as template DNA concentration, polymerase concentration, concentrations of primers and dNTPs, number of amplification cycles, denaturation and annealing temperatures, ionic strength, pH, etc. [50]. In this study [47], we found that the annealing temperature and the duration

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of each cycle were the key factors for obtaining strong signals for all four loci. Also, the temperature and time for both denaturation and extension were important factors. Almost all DNA typing and DNA diagnosis need appropriate size standards to help interpret the results. It is well known that migration times alone cannot be used for identifying DNA fragments in CE, since one capillary in an array can be very different from another [45]. Calibration based on size standards is not entirely satisfactory because base composition can influence mobility. Therefore, calibration for DNA allele typing here is based on the principle of coinjection of an allelic ladder as the absolute standard [29]. No peak broadening due to the consecutive injections or the washing step between the two injections was found. The final results are shown in Figure 3. Because of the presence of an absolute standard, the sample fragments show up as an increase in intensity for the specific allelic peaks. However, the 115bp HIV gag fragment was recognized by coinjection with a 100-bp DNA standard ladder (Fig. 4), since an appropriate standard was not available. Because there were numerous species in the PCR mixture, careful washing was necessary to clean the capillary, especially its surface. We found that it was important to wash the capillary with 0.1 M NaOH at high temperature (95°C) for 3 min, followed by washing with 5 mL MeOH and deionized water at room temperature. Five consecutive reactions were processed in this manner in one capillary. No obvious degradation or cross-contamination was observed. No purification of the PCR products was needed in most cases because the interference from dye-labeled primers or signals caused by intercalating dyes do not overlap with the DNA region of interest. Also, PCR products prepared as described can be directly injected without sample desalting. Another version of an online system for PCR analysis and for cycle sequencing was also reported recently [51]. However, here we have a much more difficult situation in the simultaneous amplification of four loci, particularly ones that readily show slippage or rehybridization, and the incorporation of an absolute DNA standard (allelic ladder) for calibration during each CE run. The elimination of a cleanup step also allows the use of low-pressure syringe pumps rather than a liquid chromatography pump.

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Figure 3 On-line PCR-CE for CTTv typing. The upper electropherogram shows the online injection and separation of CTTv standard allelic ladder only. The lower one shows the result of on-line PCR, coinjection of standard allelic ladder and PCR products, and CE separation. Notice the change of relative peak intensities in the peaks marked with (*). The genotype is named by the number of repeat sequence contained in the allele. The results are: vWA— 16, 16; THO1—9.3, 9.3; TPOX—8, 9; and CSF1PO—9, 10. (From Ref. 47.)

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Figure 4 Online PCR-CE for HIV diagnosis. The upper electropherogram shows the negative control DNA after amplification. The lower one shows the positive HIV control DNA after amplification. The 115-bp fragment of HIV1 gag is clearly identified as the peak immediately after the 100-bp fragment in the standard-size ladder. (From Ref. 47.)

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It is well known that DNA extraction from real samples (biological fluids) is always a time-consuming procedure. This procedure usually takes at least half of the total analytical time. It will be important to extend the present system to include DNA extraction and purification from real samples in an online instrument to save time, reduce labor, and minimize the exposure of workers to infectious samples. In fact, we have shown that direct PCR from human blood can be performed online in capillary tubes. We modified the approach reported by Panaccio and Lew [52]; 7.5 mg EDTA (as anticoagulant) was added to 5 mL whole blood sample. After mixing thoroughly, the blood sample was stored at 4°C until needed. One microliter of whole blood was pipetted into 20 µL pure formamide. After mixing, the vial was cooked at 95°C for 5 min. This formamide-diluted blood sample is used directly as the template in PCR. It can be stored at room temperature for several months. The PCR mixture contains 1.0 µL 10 ⫻ polymerase reaction buffer (500 mM KCl, 100 mM Tris-HCl, pH 9.0, 1% Triton X-100), 1.0 µL dNTP (2.5 mM), 1.9 µL template, 0.5 µL BSA (2.5 mg/mL), 0.5 µL MgCl2 (40 mg/mL stored in 2.5 mg/mL BSA

Figure 5 Direct online genotyping from blood. The peaks correspond to a different genotype from that in Figure 3.

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solution), 3.7 µL sterilized nuclease free water, 0.4 µL Tth DNA polymerase (5 U/mL), and 1.0 µL primers. The total volume is 10 µL. This mixture is loaded into a 40-cm-long, 360-µm-OD, 255-µm-ID fusedsilica capillary. Both ends of the capillary were sealed. PCR was then performed into a rapid-air thermocycler by the following protocol: 80°C for 1 min, then 35 cycles consisting of denaturation at 80°C for 1 sec, annealing at 40–45°C for 1 min, and extension at 60°C for 1 min. The result of simultaneous genotyping of the four CTTv loci directly from blood is shown in Figure 5. All four loci are clearly depicted. To achieve the simultaneous amplification, the experimental conditions are critical and are much more stringent than when only one locus is probed [52]. Competition among the four primers exists, so the reagent concentrations and the PCR times have to be optimized to level the competition. The conditions are also different from those used in standard thermocyclers or even the hot-air thermal cycler with glass capillaries. 5.4 MINIATURIZATION OF CE INTERFACE As discussed above, even though the sample requirement of capillary electrophoresis is in the 10-nL range, present approaches to sample injection are not compatible with submicroliter volumes. This is because physically the capillary, which is 150 µm or 350 µm in diameter, must be inserted into the sample tube to contact the sample solution. It is also necessary for an electrode to be present to implement electrokinetic injection. Sufficient liquid must be available to guarantee full contact. It may be possible to employ pressure injection [53] to eliminate the volume occupied by the electrode. Actually one would expect that pressure injection will avoid any electroinjection bias [54] that discriminates against the larger DNA fragments and the problems of electroinjection from high-salt samples. Pressure injection also avoids cross-contamination of the sample from the electrodes. Another approach is to provide electrical contact as a coating on the outside of the capillary tube. In either case, eliminating the extra electrode only reduces the volume needed for injection by a factor of 2 to 4. In analogy

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to inkjet printers, it is in fact possible to inject nanoliter volumes of sample into a capillary tube. However, much larger volumes are still needed to fill the jet reservoir before injection can be performed. It should be possible to use a miniaturized cross to inject a 10nL sample into the separation capillary. This can be modeled after K in Figure 1. If the capillary ID and the junction channel diameter are 50 µm throughout, the sample plug, as transported by gas pressure, can be 1 mm long to maintain a volume of 10 nL. The trick is to synchronize the application of the injection high voltage with the arrival of the sample plug. In Figure 1 there is already a second laser detector PMT-c to monitor the effluent from the purification column. The closer this second laser is to the injection junction, the more accurate the synchronization will be. There are no inherent problems locating the sample plug to the nearest 1 mm for injection. In fact, for multiplexed sample preparation feeding into a multiple capillary electrophoresis system, one can utilize a second CCD camera to monitor and to inject each channel independently. The injection junction can be held at a common voltage while the voltages at the exit ends of the separation capillaries can be switched independently to electrokinetically inject each sample. A new generation of miniature power supplies for photomultiplier tubes are now commercially available. The individual low-current, highvoltage power supplies can be built from such designs, and can be easily controlled by computer by using commercially available components. Our sample injection scheme is similar to cross-injection junctions which have been integrated to the separation channels in microfabricated devices [55,56]. However, there are some important differences. First, we need to inject at a high temperature to avoid rehybridization of the DNA during the transfer process. Separation, however, is performed at lower temperatures. Second, pressure is used to control liquid flow. This is necessitated by the presence of a purification column. Also, the various solutions that have to be employed to purify, rinse, run electrophoresis, and clean the junction will cause the nature of the surface to change constantly over the course of the operation. Electro-osmotic control of the flow is simply not possible. Third, the junction(s) will still need to be connected to the purification column,

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separation capillary, waste, and the cleaning solutions. Since the separation column will have to be flushed out and filled with polymer matrix at moderate pressures, special bonding processes will need to be developed to maintain the integrity of the microfabricated device. Fourth, even though separation of the Sanger fragments can be performed in microchannels, a certain minimum length is needed to resolve the larger fragments in electrophoresis. Microfabrication of an integrated device therefore does not provide any overall size advantage. On the other hand, this does not mean that a standalone junction region should not be microfabricated, especially when large numbers of junctions are eventually used in parallel. 5.5 MINIATURIZATION OF SANGER REACTION Our demonstration of online, one-step sequencing from template to called bases provides the basis for miniaturization of these front-end operations. Everything should therefore be performed in 50- to 100µm-ID capillary tubes to achieve complete integration at the 10- to 100-nL scale. Since present technology (e.g., robotic work stations) typically requires the use of sample volumes of 1 µL, and since standard technology calls for 10-µL sample volumes, our anticipated cost savings related to templates, reagents, dye labels, buffers, separation matrix, etc. is 100- to 1000-fold. The first demonstration system [14] was not designed to handle 10-nL samples. First, we needed to produce sufficient amounts of products to directly evaluate the performance by using a standard sequencing instrument. Those instruments require loading 1-4 µL of sample into each well. Second, a larger sample volume permitted the creation of longer plugs of samples within the column. This makes sample transport and synchronization of the various steps in the process much easier to control in a manual operation. Third, our template was premixed with the enzyme and the dye-labeled terminators prior to introduction into the capillary system. This ‘‘injection’’ step therefore started with samples in the range of a few microliters although much less was needed. Fourth, the connections and valves are components taken from routine liquid chromatographic (LC) instrumentation. There, ‘‘dead

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volumes’’ are in the order of a few microliters. Fifth, the commercial air thermal-cycler was designed for relatively long and large-ID capillary tubes. All of the above are features that can be readily modified for operation at the 10-nL range. First, the successful development of a miniaturized junction, as described above, will allow the direct analysis of the products by capillary electrophoresis. Our successful initial study [14] shows that it is no longer necessary to use a standard DNA sequencer to check the various subsystems. Second, computer automation and the use of a constant-pressure source to drive the fluidics means that delivery of sample plugs to the nearest 1 mm (10 nL equivalent volume) should be feasible. The analogy here is flow-injection analysis, which involves much larger volumes only because the diameter of the tubing is much larger. Pressure-driven Pouiselle flow is highly reliable (as opposed to electro-osmotic flow, and has already been implemented in commercial CE instruments for injecting 10-nL samples. Third, one can envision the template being injected by pressure into a capillary tube at the 10-nL level. This plug is first transported to a cross junction, similar to that described above. The inline junction goes to the reaction capillary. The two extra inlets will contain enzyme and dye labels, respectively. With independent control of the pressures and the ID of the inlets, well-defined mixing ratios can be achieved. Even if the plugs of template, enzyme, and dye labels are not well mixed at this junction, the fact that cycle sequencing is operated at elevated temperatures means that complete mixing can be achieved just by axial diffusion. We note that the diffusion distances involved if three plugs are injected consecutively will be only 3 mm. This arrangement will essentially keep the enzymes and dye labels as stock reagents delivering only as needed to the cross junction. There is thus a true savings in reagent costs. Fourth, one needs to modify the valves to become on-column controls, as described below. There is no connection and therefore no extra dead volume. Fifth, conditions will have to be developed to allow cycle sequencing inside 50- to 75-µm-ID capillaries. We have already shown that the fused-silica surface can be pacified if necessary with added BSA [14,47]. Reduction of the column diameter affects heat capacity. So, hysteresis in the cycling timing and cycling temperatures will be different compared to before.

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5.6 MINIATURIZATION OF THE PURIFICATION COLUMN Previously, we used 1-mm-ID PEEK tubing to pack our own purification columns. The elusion plug in liquid chromatography has a volume that is roughly proportional to the cross-sectional area of the column. By going from 1-mm ID to 100-µm ID, the plug is reduced in volume by 100-fold. We note that 50-µm-ID packed capillaries are already available commercially for microscale LC applications. The purpose of the purification column is to isolate the labeled Sanger fragments from the unreacted dye labels. Such does not demand very high chromatographic efficiencies. It should be possible to pack 100-µm-ID columns with Sephadex-25 to produce the targeted reduction in volume. The use of smaller-ID columns also reduces the costs associated with the packing material and the elusion buffer. A second issue in miniaturization is the back pressure that is needed to generate the same linear velocity. On first inspection, Pouiselle law will predict that the pressure will go up as the column dimensions are reduced. However, this is more complicated since the main source of back pressure is the restricted channel spacing between the packed particles, not the ID of the support tube. Theoretical plates in separation also refer to channel dimensions and not the ID of the tube housing. One does not need to substantially reduce the particle size of the Sephadex packing. So, there should be little change in back pressure or in separation efficiency on miniaturization. Only the column capacity is decreased, but then there will be much less sample. Eventually, the ID of the capillary tube will contribute, but not at the 100-pm range. A third issue is dilution of the sample plug after passing through the purification column. This is a natural effect of retention during separation. There will be dilution, but our studies have already shown that the factor of 10–50 dilution does not result in a factor of 10–50 reduction in S/N due to electrofocusing at the injection tip. The S/N obtained is more than sufficient for good base calling. Actually, the dilution effect likely prevents the DNA fragments from rehybridizing with each other and from forming secondary structures. This is the primary reason why additional denaturant was not needed in the system. If necessary, it is possible to utilize more of the (longer) diluted sample plug by

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performing electroinjection while (pressure) flowing the useful portion of the effluent from the purification column across the junction. The Sanger fragments can therefore be reconcentrated at the tip of the separation capillary for electrophoresis [54]. For this, we note that ‘‘normal’’ injection times in CGE sequencing are 30–45 sec at the running voltage. No noticeable band broadening was observed. Since the separated plugs are monitored while they elute from the purification column, electroinjection during flow can be implemented under computer control. 5.7

MULTIPLEXED ONLINE REACTION AND ELECTROPHORESIS FOR DNA SEQUENCING

The above efforts to miniaturize the various front-end steps in DNA sequencing would have little value if the process can only be performed one at a time. This would then still be incompatible with capillary array electrophoresis or microchannel electrophoresis. Fortunately, the above concepts are all inherently suitable for large-scale multiplexing. It is this total integration of front-end manipulation to the mature technology of multiplexed CE that brings true cost savings while preserving the high-speed, high-throughput advantages. The first issue is automation of the apparently complicated valving system shown in Figure 1. While further research may allow one to reduce the number of cleaning steps, even the present arrangement is ready for automation and multiplexing. The switching valves A through E can be replaced by commercially available computercontrolled switching valves. The glass syringes (S in Fig. 1) can be replaced with simple, stepping-motor-driven syringes identical to those already used for the cross region (P in Fig. 1). The important point is that only one channel each of S, P, or W is needed even for a large number of parallel capillary reactions. What one needs is a distribution network to feed into each channel, since the protocol is identical in each. For example, the flow in each capillary is expected to be in the order of 1 µL/min at the maximum. For 1000 parallel samples, total flows are still around 1 mL/min, which is typical of HPLC flow rates. For the purification step, the original design involves flow rates of 10 µL/min. However, the ID of the purification column will be reduced

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eventually, thereby maintaining the same total flow as in typical HPLC instrumentation. As for pressure, 1000 75-µm-ID capillaries have a cross-section roughly equal to a 2.2-mm-ID open tube. The back pressure does not go up on multiplexing. The syringes will only need to be of large enough ID to provide enough total solution. The only high-pressure component is the purification column. We found that the back pressure developed is only around 200 psi per capillary. Again, the back pressure does not go up on having more columns. The ID of the HPLC pump used is clearly sufficient for 1000 capillaries. A curious situation exists if the back pressures of the individual channels are not identical. Flow will favor the channels with the least resistance (back pressure). As long as the back pressures are within a reasonable range, some flow will pass in each channel. To this end, automation of the offline column packing process to produce fairly even back pressures will be important. This is simply repeating current manufacturing protocols for commercial HPLC columns, where the back pressures are substantially higher than here. As discussed above, the arrival of the purified Sanger products can be independently monitored to initiate electrokinetic injection. There is no need for the sequencing data to be synchronous in the capillary array. The third issue is the reduction of the number of valves used, even though each valve (A though E in Fig. 1) can be computer controlled. The system can be modified by using reversible syringe pumps. The idea is shown in Figure 6. For each sample channel, three onoff on-column valves (V1 to V3) will suffice for Sanger reaction and purification. With V1 blocked and V2 and V3 open, the template plus reagent plug (previously mixed in a cross) can be transferred from below to the center of the reaction capillary loop inside the hot-air thermal cycler. V2 and V3 are then blocked during cycle sequencing. This essentially pressurizes the sample plug, so there will not be boiling or segmentation. After reaction, V2 and V3 are opened while V1 remains blocked to bring the sample plug up to the T junction. V2 and V3 are then blocked while V1 is opened while the syringe pump reverses to send the product mixture to the purification column. Elution continues to put the correct separated fraction at the injection cross. Fluorescence monitored via a split portion of the same 514-nm excitation laser syn-

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Figure 6 Schematic of the complete multiplexed and integrated instrumental design with eight capillaries. Stars at I, U1, and U2 represent the multiplexed freeze/thaw valves. The T assembly is made up of eight pieces of commercial junctions stacked together. These connect to the manifold M1, the SEC purification columns, and the reaction loops. The cross-assembly is made up of eight pieces of standard crosses packed together with built-in heaters. V8 is an eight-position motorized titanium valve with a center port. S1 is a two-position motorized PEEK valve. V6 is a six-position motorized PEEK valve. (From Ref. 57.)

chronizes the individual injection into the capillary array. Later, with V1 blocked and V2 and V3 open, the reaction loop can be cleaned. Cleaning solutions and new separation matrix can be introduced to the separation capillary at this time. Then, with V2 and V3 closed and V1 open, the purification column can be regenerated. The entire operation is then reduced to only two multiposition valves. In fact, Figure 6 shows how multiplexing can be implemented [57]. Two distribution manifolds connect all the channels. This is possi-

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ble because all channels are treated in the same way. Yet, the samples are always isolated from each other through reaction, purification, injection, separation, and detection. Not shown are multiplexed automated filling, cleaning, and regeneration of the separation columns. This is already dealt with in the commercial versions of the capillary array electrophoresis systems [58]. The final issue is that each channel must have its own set of oncolumn on-off valves. Since V2 and V3 always open and close at the same time, only two sets of valves are required in practice. The freezethaw valving system developed for CO2 gas jets [59] for liquid N2 operation. Figure 7 shows the basic concept. The independent capillary channels are bundled together to intersect the flow path of cold N2 gas from a liquid nitrogen container. Freezing is initiated by solenoid valves which control liquid N2 flow, much like what is done in cryogenic vacuum technology. To facilitate thawing after the N2 flow is stopped, a local heating element is turned on. A temperature probe

Figure 7 Multiplexed on-column freeze-thaw valves. Liquid nitrogen is controlled by a gas solenoid to freeze the solution. A local heater is used to thaw the solution. A thermistor provides feedback to the controlling computer to indicate the status of the valves. (From Ref. 57.)

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serves as feedback for computer control. The entire assembly is wrapped in insulating material to increase efficiency. Such a system controlled by commercial software such as Labview can produce freezing and thawing in a few seconds. As explained before [59], the frozen buffer solution blocks all electrical current and can withstand quite high pressures. The local heat capacity is quite small to allow multiplexing of many capillaries. We note that 100 capillaries laid side by side only occupy a width of 1.5 cm, which easily fit into one multiplexed valve. Based on these concepts, an integrated and multiplexed online instrument starting from DNA templates to their primary sequences has been demonstrated [57]. The instrument automatically processes eight templates through reaction, purification, denaturation, preconcentration, injection, separation, and detection in a parallel fashion. A multiplexed freeze/thaw switching principle and a distribution network were utilized to manage flow and sample transportation. Dye-labeled terminator cycle-sequencing reactions are performed in an eight-capillary array in a hot-air thermal cycler. Subsequently, the sequencing ladders are directly loaded into separate size exclusion chromatographic columns operated at ⬃60°C for purification. Online denaturation and stacking injection for capillary electrophoresis are simultaneously accomplished at a cross assembly set at ⬃70°C. Not only the separation capillary array but also the reaction capillary array and purification columns can be regenerated after every run. The raw data allow base calling up to 460 bp with accuracy of 98%. The system is scalable to a 96-capillary array and will benefit not only high-speed, highthroughput DNA sequencing but also genetic typing. The instrumental operation can be divided into four steps, which are loading of cycle sequencing reaction mixture into the microreactor array, loading of these reaction products into the SEC purification columns, heart cut injection into multiplexed capillary array, and sizebased separation and detection of the sequencing ladder. Figure 8 shows the key steps to load samples separately into the reaction loops and the purification columns before and after the Sanger reactions. With valve I closed and valve U opened, 22 µL was aspirated into each reaction capillary from the microtiter plate where a row of premixed templates and reagents were placed. Then, another plug of 8.5 µL deionized water was loaded into each capillary. This placed the

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Figure 8 Online operation protocol from reaction capillary to purification column. Open star means MFTV is on. Filled star means MFTV is off. Wave filling represents the buffer or water. Dot filling denotes the reaction mixture. (A) Simultaneous reaction and regeneration of SEC column. During cycle sequencing reaction, the 1 ⫻ TE buffer flushes the purification column with valve U off. (B) Loading. When reaction is complete, the syringe pump aspirates the reaction products above the T assembly with valve I off. (C) Micropump drives the Sanger products into the purification columns at 20 µL/min/ch with U closed. (From Ref. 57.)

reaction plug in each capillary right inside the thermal cycler. After opening valve I and closing valve U, the cycle sequencing reactions were started (Fig. 8A). During reaction, 1 ⫻ TE buffer continuously flushed the purification columns at a flow rate of 20 µL/min/ch (/ch means per channel) from the micropump. After reaction, valve I was closed and valve U was opened, so that 22.5 µL/ch was pulled into the Teflon tubing region above the T assembly (Fig. 8B). After switching off valve U and switching on valve I, reversing the flow direction drove these plugs into the purification columns (Fig. 8C). Purification was performed at a flow rate of 20 µL/min/ch with 1 ⫻ TE buffer at 60°C as controlled by the hot water bath.

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Once the purification step started (by selecting the micropump at valve S1), there were two operations necessary to prepare the crossassembly junction for injection. The temperature at the cross-assembly was set at 70°C while deionized water flowed over the injection junction at 200 µL/min/ch from the syringe pump. The continuous transverse flow of water at the injection end of the separation capillaries causes dialysis at the interface of the sieving matrix to create a low ionic strength zone. This promotes stacking injection for pre-concentration of the analytes. When the Sanger fragments reached the LIF detector at point A, as recognized by the onset of a peak, electrokinetic injection to the separation capillaries was carried out at 150 V/cm for 2 min. When injection is completed at about one-third of the peak height on its falling edge, valve I was turned off and valve S1 was also switched to the syringe pump position. Buffer 1 mL of 1 ⫻ TBE was used to clean out the deionized water and reionize the injection zone at a very low flow rate to avoid losing the injected fragments. The 1 ⫻ TBE transverse flow was then programmed to 500 µL/min/ch during electrophoretic separation at 150 V/cm. More importantly, the temperature at the injection cross-assembly needs to be lowered to room temperature as quickly as possible after restoring the ionic strength at the cross-junction. The MFTV valve I can be turned on and regeneration of the purification columns and reaction capillaries can proceed after the electrophoresis current has stabilized (about 40 min). The microreactor array, the cross-assembly and the T assembly were simply regenerated by flushing with 0.2 M NaOH solution, 1 ⫻ TE buffer, methanol, and deionized water sequentially. The purification columns can be recovered by NaOH solution and 1 ⫻ TE buffer as described previously. Achieving uniform flow rates in each channel is very critical to success of this multiplexed operation. The accuracy of sample loading, the timing of ‘‘heart-cut’’ injection at the cross-junction, the similarity of transverse flow at the interface between the purification columns to the separation capillaries all depend on how uniform the flow rates are among the channels. Here, the capillary ID, the pressure drop, temperature, and viscosity among the channels are essentially identical. Therefore, the capillary length is the most important parameter toward achieving a uniform and controllable flow rate.

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In the microreactor array, the variation in flow rates can be experimentally determined by the difference in aspirated lengths of liquid plugs viewed outside the capillaries. We found there is ⬃2% relative standard deviation among the channels. So, 20 µL of reaction mixture occupies 40 ⫾ 2.4 cm of capillary, the required distance between the two U valves. A distance of 7 cm was thus introduced between the MFTV valves U. For the same reason, 30 µL of Teflon tubing between the manifold M1 and the T assembly was used to guarantee independent sample manipulation in the eight channels. Although 22.5 µL of liquid plug was loaded into this Teflon tube region, only 15 µL of sequencing products was injected into the purification column. The other 7 µL of deionized water was also loaded into the purification column but there are no observable negative effects in terms of purification and regeneration. In the purification columns, the main source of back pressure is the restricted channel spacing between the packed particles, not the ID. If the back pressure of the individual channels are not identical, flow will favor the channels with the least flow resistance. Moreover, the arrival time can also be adjusted by manipulating the capillary length after the purification column. It is clear that a synchronous ‘‘heartcut’’ injection can be performed with careful engineering and relatively uniform column packing. Once the system has been calibrated, only one of the purification columns needs to be monitored. The electropherograms (raw data) at the blue channel from a completely multiplexed and online run starting from M13mp18 templates are shown in Figure 9. We found variations of intensities among capillaries. This fluctuation is caused by heterogeneity in the efficiencies of individual sequencing reactions, efficiencies of the stacking injection, and temporal variations in the ‘‘heart-cut’’ injection. All electropherograms, however, have very high S/N ratios. The S/N of the blue channel for capillary 8 is the lowest, but is still 39 at 453 bases, which is clearly enough to call bases. The migration time and resolution of DNA fragments in each capillary were not completely uniform among the capillaries. This is probably attributable to inhomogeneities in the gel matrix, variations in capillary length, and reproducibility of the (separation) capillary regeneration and coating procedures. Naturally, in DNA sequencing,

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each channel is an independent experiment and quantitative reproducibility is not required. For all eight capillaries in Figure 9, sequences can be called based on the raw data up to 400 bases with an accuracy of 98%. The best capillary produced this same level of accuracy out to 460 bases (at 120 min). The ability to recondition the multiplexed system is a major issue. We were able to use the same microreactor array and the same set of purification columns over a period of 4 months for ⬎20 runs. We have not observed any significant degradation in performance. Although the reaction efficiency may vary from run to run and from capillary to capillary, it is not critical for DNA sequencing as long as it provides sufficient sequencing products to maintain a minimum S/N. The turnaround time for this demonstration is about 5.5 hr including 2.5 hr for separation, 2.5 hr for reaction, and 0.5 hr for purification and sample loading. The regeneration of the reaction capillaries can be done during separation and vice versa. With a slight modification of the protocol described here, one should be able to carry out Sanger reaction and electrophoretic separation at the same time in the two subunits of the system. Furthermore, the reaction time in the air thermal cycler can be as short as 25 min [4]. Reducing the separation time to 1 hr with 1000 bases sequenced has also been reported recently [60]. Producing 1.2 million bases of sequencing data per day per instrument should therefore be achievable if the system is scaled up to 96 channels. Figure 9 Electropherograms showing simultaneous sequencing of eight M13mp18 DNA templates with the online integrated and multiplexed system starting from templates. Reaction plug: 40 cm out of 77 cm of 250-µm-ID and 360-µm-OD fused-silica capillaries; flow rate, 20 µL/min/ch; purification: Sephadex G-25-50 columns set at 60°C; injection: ‘‘heart-cut’’ injection at 70°C, with ⬃2 mL of water preconditioning; separation: 60 cm effective length of bare capillary filled with 1.5% high-molecular-weight PEO and 1.4% low-molecular-weight mixture, 150 V/cm; detection: CCD camera; excitation: 15 mW 514.5 nm Ar ion laser with exposure time of 300 msec. Capillaries in the array are labeled from 1 to 8 according to proximity to the excitation laser. (From Ref. 57.)

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MULTIPLEXED ONLINE PCR ANALYSIS AND GENOTYPING FROM BLOOD

An automated and integrated system for DNA typing directly from blood samples has also been developed [61]. The multiplexed eightarray system is based on capillary microfluidics and capillary array electrophoresis. Three STR loci, vWA, THO1 and TPOX, are coamplified simultaneously in a fused-silica capillary by a hot-air thermocycler. Blood is directly used as the sample for polymerase chain reaction without any pretreatment. Modifications of standard are necessary for direct PCR from blood. A programmable syringe pump and a set of multiplexed liquid nitrogen freeze/thaw switching valves are employed for liquid handling in the fluid distribution network. The system fully integrates sample loading, PCR, addition of an absolute standard, online injection of sample and standards, separation, and detection. The genotypes from blood samples can be clearly identified in eight parallel channels when the electropherograms are compared with that of the standard allelic ladder by itself. Regeneration and cleaning of the entire system prior to subsequent runs are also integrated into the instrument. The instrumentation is compatible with future expansion to hundreds of capillaries to achieve even higher throughput. Because there was no need for purification of the PCR products, all connections between the T assembly and the cross-assembly were open tubes in this study, as shown in Figure 10. One syringe pump could provide enough pressure for liquid transfer in even 1000 channels. No expensive high-pressure pump was necessary, in contrast to DNA sequencing where a higher pressure is needed for sample cleanup [57]. Also, only one multiposition valve was needed for the entire operation. For multiplexed operation, flow uniformity was critical because all operations, especially sample injection, were simultaneous for all channels. The flow rates in the eight channels here were quite uniform because of the open tubular character. The capillary length was the determining factor because all other parameters such as the diameter of capillary, solution viscosity, and temperature were essentially identical in each channel. One only needed to keep the capillary and the Teflon tubing lengths identical in each channel. Also, every port of the T assembly and the cross-assembly should be kept identical. The flow

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Figure 10 Schematic of the complete multiplexed and integrated instrumental design for PCR analysis with eight channels. Different functional capillaries were connected by a T assembly and a cross-assembly. Stars represent the freeze/thaw valves V1 and V2. A syringe pump and a six-way selection valve were used to load and distribute liquids for various purposes. A CCD camera was used to collect fluorescence from the eight capillaries simultaneously. The temperature control unit at the cross-assembly was used to denature DNA prior to injection. (From Ref. 61.)

uniformity was verified by observation with a CCD camera above the transfer capillary bundle near the cross-assembly. Figure 11 shows the individual steps of sample introduction, PCR reaction, standard ladder loading, and injection. To put the PCR mixture and the allelic ladder in the appropriate regions of the capillary tubes, the volumes of aspiration and dispensing from the syringe must be calculated from the capillary length and ID. The time delay for initiating electrokinetic injection (13 sec) was calculated from the syringe

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Figure 11 Online operational protocol for sample introduction, PCR reaction, standard ladder loading, and injection. The operation was controlled by two freeze/thaw valves and a reversible syringe pump. (From Ref. 61.)

flow rate and the transfer capillary volume. The injection duration (33 sec) was obtained by dividing the total volume (28 µL) of the PCR mixture and the standard ladder solution by the flow rate of the syringe pump (50 µL/ch/min). The actual liquid handling performance was found to be very accurate based on these calculations. Each channel was always filled with H2 O or other solutions during the whole process to prevent bubble formation at the cross-junction. The standard ladder was loaded as a plug following the plug of PCR mixture. The two plugs were pushed to the cross-junction, then injected one after the other. This method simplified the instrument design and was efficient because no additional channels were attached for standard loading. Repeated injection of the standard ladder over many runs from the same set of vials was possible. This reduces the amount and thus

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the cost of standard solutions needed. Previous studies [31,62] showed that coinjection of PCR products and the allelic ladder (as an absolute standard) provides reliable identification of unknown alleles over a wide range of relative peak intensities. An intensity ratio between 0.5 and 1.5 is recommended [62]. This can be achieved by adjusting the volume of the solution plug or the concentration of the allelic ladder solution. The capillary reactors must be cleaned carefully after PCR and allelic ladder loading. An extremely small amount of the allelic ladder or the PCR products can be amplified easily by PCR during the subsequent run to cause interference. The washing step was evaluated by performing negative-control PCR, in which water was used instead of blood while all other reagents were kept identical. No DNA fragments were found except for the primer peaks. The ability to reuse the array of reaction capillaries is vital to the ruggedness of an online system. After all, it is not just how fast or how many samples one can handle in one run, but also whether the operation can be repeated over and over again in rapid succession that is important for large-scale applications. Figure 12 shows the results of automatic genotyping from blood to readable results in the multiplexed format. The peaks before 40 min are due to residual primers and fluorescent contaminants in the samples. As seen from the electropherograms, purification before analysis was not necessary because the allele fragments are all ⬎100 bp. Direct PCR from blood often shows small contamination peaks especially in the vWA region. However, the true allele peaks were much more intense so that highly confident allele identification was still possible. The baseline fluctuations were more severe in channel 3 and 8. This may be related to unevenness in manual gel-filling and chemical noise in this run. The peak intensities of electropherograms also varied among the different channels even for identical blood samples. This is likely due to the variability of PCR efficiency, stacking injection efficiency, and nonuniform detection characteristics across the channels. However, such variations will not affect the identification of genotype as long as the PCR, injection, and detection provide enough signal to alter the relative peak intensities in each allelic region in our absolute standard method [29,62]. In this sense, the PCR specificity is more important than its efficiency.

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Figure 12 Electropherograms showing simultaneous genotyping in eight channels from blood to final results. Injection: temperature 90°C, voltage 150 V/cm, time 33 sec. Separation: voltage 150 V/cm, effective length 60 cm, matrix 1.5% Mn 8,000,000 and 1.45% Mn 600,000 PEO solution. Laser: 15 mw 488 nm argon-ion laser. Detection: CCD through a 520-nm long-pass filter, exposure time 300 msec. Capillaries in the array are labeled from 1 to 8 according to proximity to the excitation laser. Two blood samples were analyzed. Channels 1 to 4 show sample 1 and channels 5 to 8 show sample 2. The lower electropherogram is the vTT allelic ladder by itself. The genotype (asterisks) was identified by an increase in the relative intensities of the peaks within each locus when the sample is present. (From Ref. 61.)

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The total time for one run is 2.5 hr. However, since cleaning and regeneration of the separation capillaries can be implemented during the PCR reaction and since a subsequent set of PCR reactions can be run during electrophoretic separation of the first set of samples, the real cycle time can be shortened to 1.3 hr. We note that because of the concept of absolute standards [29,62], these results are obtained for one-color detection of the DNA fragments. Reoptimization of the PCR conditions should allow two or more sets (different loci) of multiplex PCR to be performed simultaneously by using two or more sets of differently labeled primers and multicolor detection. If so, forensic DNA fingerprinting at a crime scene can be achieved in 2.5 hr with identification at errors rates ⬍1 in 108, which is the current legal definition of a unique match. Full integration and automation of Sanger reaction for DNA sequencing and direct PCR from blood were achieved here in capillary tubes. One can envision transferring this protocol to microfabricated devices [63–65] or other solid-phase microreactors [66]. Capillaries are more compatible with pressure flow for gel filling, for avoiding irreproducibility inherent to electroosmotic flow and for sample purification in chromatographic columns [57]. Capillaries are also more flexible during the research and development stage because minor modifications in the hardware do not require completely rebuilding the system. On the other hand, microfabricated devices do not require liquid valves (substituting instead multiple high-voltage relays), can be made more compact and are more adaptable to mass production to reduce cost once the optimal protocol is established. Our future efforts will focus on incorporating larger numbers of channels and miniaturization of the reactor volume by reducing the capillary diameter to further lower the cost of reagents. Applications can be expanded using this concept to other sample preparation protocols prior to CE, such as drug screening and peptide mapping. ACKNOWLEDGMENTS The author thanks the many coworkers in his laboratory who have contributed to this work, especially H. Tan and N. Zhang. The Ames Laboratory is operated for the U.S. Department of Energy by Iowa State

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University under contract No. W-7405-Eng-82. This work was supported by the Director of Science, Office of Biological and Environmental Research, and by the National Institutes of Health.

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63. Woolley, A. T., Hadley, D., Landre, P., deMello, A. J., Mathies, R. A., and Northrup, M. A. Anal. Chem. 1996, 68, 4081–4086. 64. Woolley, A. T., Sensabaugh, G. F., and Mathies, R. A. Anal. Chem. 1997, 69, 2181–2186. 65. Waters, L. C., Jacobson, S. C., Kroutchinina, N., Khandurina, J., Foote, R. S., and Ramsey, J. M. Anal. Chem. 1998, 70, 158–162. 66. Soper, S. A., Williams, D. C., Xu, Y., Lassiter, S. J., Zhang, Y., Ford, S. M., and Bruch, R. C. Anal. Chem. 1998, 70, 4036–4043.

6 Interfacing Capillaries to Microseparation Devices for Sample Introduction Steven W. Suljak and Andrew G. Ewing Pennsylvania State University, University Park, Pennsylvania

6.1 INTRODUCTION Capillary zone electrophoresis (CZE) has become a high-efficiency separation method ideal for small-volume samples [1]. CZE has been widely used to separate material taken from complex biological microenvironments such as single cells [2–6]. It has also been applied to high-resolution DNA separations through the use of gel-filled capillaries [7]. Since the early 1990s, the drive toward miniaturization has led to the development of capillary electrophoresis on microfabricated devices, frequently referred to as chips [8–10]. These devices have integrated sample and buffer reservoirs and fluidic channels onto a single substrate (generally glass or a polymeric material), and have retained the high resolving power of capillary electrophoresis. The miniaturization of the separation channel has allowed the application of electric fields as high as 2500 V/cm, resulting in high efficiencies in very short times. Such separations have generated up to 18,600 theoretical plates per second [11]. Sample introduction onto chips, however, has proven to be troublesome. The most common injection technique has incorporated a T129

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style injector in which sample is electrokinetically loaded perpendicularly across the separation lane (Fig. 1). Potential is then applied to the separation channel, and the sample plug is separated into its components. Even though this method has the ability to separate picoliter volumes, significantly more sample is needed to fill the sample reservoir and lane. Also, high throughput of multiple samples requires a separate injection reservoir or the use of different separation channels for each sample. Additionally, monitoring dynamic processes has proven to be difficult on chips since the technique is inherently incremental, incorporating a brief sampling period followed by electrophoretic separation. To circumvent this difficulty, fast, repetitive CZE separations have been performed on chips [12,13], and separations as fast as 0.8 msec have been reported [14]. However, with these methods material is still sampled and separated in discrete intervals. An alternative approach to introducing sample onto a microfabricated structure involves the use of a fused-silica capillary to inject sample into an open rectangular channel [15–19]. The use of a capillary to deliver sample to the separation channel has significant advantages over the methods currently used in other microfabricated systems. Injecting analytes with a capillary permits direct sampling from the environment of interest. The small size of the capillary facilitates sampling

Figure 1 Photograph of a sample plug (o-phtaldialdehyde-labeled phenylalanine and arginine) formed in a ‘‘double-T’’ injector. Voltages are applied only to the sample and sample waste reservoirs to position a plug of sample within the separation chamber before application of the separation potential. (From Ref. 10.)

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from low volumes or localized areas (such as the exterior of a single cell). Multiple samples can be injected easily by moving the capillary from one sample to the next. Finally, the use of capillary sample introduction allows the continuous transfer of a dynamic sample to the separation device. The coupling of the sampling capillary to a rectangular channel, termed channel electrophoresis, allows either continuous deposition of a dynamic sample or the sequential deposition of multiple samples across the channel width. This chapter will describe the details of the system design, delineate some of the variations that have been developed to expand the potential of channel electrophoresis, and describe the various applications for which the technique has been utilized. 6.2 SYSTEM DESIGN Channel electrophoresis incorporates three basic components: a rectangular channel in which electrophoretic separation occurs, a capillary used to inject sample onto the channel, and a spatially sensitive detection scheme (Fig. 2). All three elements are crucial to achieve continuous, high-throughput separations.

Figure 2 Experimental setup of the channel electrophoresis technique. A capillary is coupled to a channel that is suspended across two buffer reservoirs. The capillary is attached to a translation stage and a stepper motor that moves the capillary across the channel entrance. Analytes separate in the channel due to the potential field applied and are detected by the spatially sensitive detector at the channel exit.

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Rectangular Channels

A key component of the channel electrophoresis system is the channel structure itself. Separations occur in a wide (centimeter range), but narrow-thickness (micrometer range) rectangular volume. The concept of incorporating a rectangular geometry into an electrophoretic device was first introduced in 1937 by Tiselius for the separation of a variety of proteins [20]. Since that time, a variety of reports have examined both theoretical and experimental advantages of using rectangular capillaries versus more traditional cylindrical types [21–26]. First, dissipation of Joule heating is more efficient in rectangular structures, assuming a large width-to-thickness ratio [20,23,25]. As the thickness is decreased, heat dissipation is especially effective, allowing the application of large electrophoretic potentials. Second, since widening the rectangular channel does not attenuate heat dissipation, a large increase in sample capacity can be gained [25,26]. This is an especially important factor in channel electrophoresis, since sample is continuously introduced into the channel. Third, rectangular structures offer additional advantages to optical detection systems. Larger path lengths can be achieved, yielding improved detection limits. At the same time, the flat surfaces of the rectangular geometry lead to less scatter and optical distortion than those resulting from cylindrical capillaries [26]. The structures used for channel electrophoresis are facilely constructed with two planar surfaces separated by a spacer that is used to determine the internal height of the structure (Fig. 3). Channels have

Figure 3 Schematic of the fabrication of a channel. Spacers are placed along the edges of two planar surfaces before adhering the surfaces together, forming an open rectangular channel (front view) in which all separations take place.

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been made with a variety of materials, but most have been fabricated using standard borosilicate glass slides or polished glass plates (quartz or borofloat). The spacers employed have included Scotch tape, borosilicate glass microspheres, and silica chromatography beads. The Scotch tape, approximately 57 µm in height, is placed along the lengths of one glass plate before adhering the second plate. Smaller-height channels (low to submicrometer sizes) can be achieved using appropriatelysize spherical particles. These particles are mixed in an epoxy and applied to the lengths of one plate. The channel is then constructed by clamping the second plate onto the first and curing the epoxy. Since the internal heights of such channels are defined only by the size of the spacers, a wide variety of channels with different sizes can be easily constructed. Filling the channels can be accomplished either before or after channel construction, depending on the viscosity of the separation matrix being used. Channels made in this fashion are convenient because they can be recycled by simply removing the adhesive and remaking the channel. 6.2.2

Sample Introduction

Sample is introduced into the rectangular channel structure via a standard fused-silica capillary. One end of the capillary is placed directly in or near the sample of interest. Application of a potential across the capillary causes electrophoretic migration of the sample to the opposite end, situated in the channel entrance. Sample is continuously deposited by attaching the capillary to a stepper motor that translates the capillary across the width of the channel entrance. A separate potential applied across the length of the channel causes analytes to separate as they migrate linearly through the channel. By continuously depositing sample across the channel entrance, a series of parallel separations are, in effect, established in the channel. A depiction of how separations occur in the channel is shown in Figure 4a. In this schematic, a 3-component mixture is continuously injected onto the channel from the sampling capillary. Initially (time 1), the capillary injects the mixture on the left side of the channel, and its components migrate linearly toward the channel exit. As the capillary progresses to the right (time 2) across the

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Figure 4 A depiction of the channel electrophoresis technique. (a) Schematic of three different stages in the sampling and separation process. The panel on the left (time 1) shows a mixture of three analytes being deposited on the channel. At a later time (time 2) the capillary has moved the right and is now depositing sample in the center of the channel, while the material introduced earlier has begun to migrate down the channel and separate. In the third panel (time 3), the material deposited on the left has completely separated, while the material at the right is just entering the channel. (b) The resulting electropherogram of the separation above is shown indicating the separation of three distinct components. The diagonal feature of the separation is due to the difference in the time of deposition by the capillary from one side of the channel to the other.

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channel entrance, the material deposited earlier has begun to separate electrophoretically into its individual analyte bands. By the time the capillary is injecting sample at the right side of the channel (time 3), the sample injected earlier on the left side has completely separated. The resulting electropherogram (Fig. 4b) exhibits three bands, one for each analyte, separated by a constant amount of time since the sample in each ‘‘lane’’ spends the same total time migrating through the channel. The diagonal nature of the bands is the result of the different times at which samples are injected across the channel. Although this schematic shows only one pass of the capillary across the channel entrance, the direction of capillary movement can be reversed and moved back and forth as many times as desired. The parallel lanes of separation established within a channel permit two different types of experiments. The capillary can be placed in a dynamic environment and used to continuously inject sample, allowing continuous, time-resolved channel separations. Multiple samples can also be injected in serial fashion into the capillary and then be deposited across the width of the channel, resulting in high-throughput separations. Both situations will be examined later in more detail. 6.2.3

Detection Methods

A variety of detection methods have been coupled to channel electrophoresis with capillary sample introduction, all sharing the crucial feature of retaining spatial selectivity. Since the location of an analyte eluting at the channel exit corresponds directly with its injection location, and thereby its time of injection, the detector must be sensitive to this position. To date, detection of channel separations has been accomplished using laser-induced fluorescence (LIF) coupled to a photodiode array [15,27,28] fluorescence with a charge coupled device (CCD) [29–37], and electrochemically with an array of platinum microelectrodes [38–42]. Fiber Optics/Photodiode Array Detection The demonstration and early characterization of channel electrophoresis have been accomplished by employing two fiber-optic arrays coupled to a photodiode array, as shown in Figure 5 [15,27]. Using this

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Figure 5 Schematic representing the setup of the laser-induced fluorescence detection scheme with fiber-optic arrays and a linear photodiode array. (From Ref. 28.)

mode of detection, an 80-fiber array is used to deliver excitation light from a HeCd laser (325 nm) to the channel exit. The laser is either directly connected to a bundled end of the array or first directed through a divergent plano convex lens. The other ends of the fibers are epoxied side by side at the channel exit, directing light through the length of the channel. This configuration allows fluorescent analytes to be observed visually as they exit the capillary, enter the channel, and begin to separate electrophoretically within the channel. Quantitative detection is achieved by collecting fluorescence emission at the channel exit with a second linear array of fiber optics. This array of 150–160 fibers is epoxied to the top channel plate, perpendicular to the channel plane, and the collected signal is directed to a linear array of 512 photodiodes such that spatial integrity is preserved. Spatial resolution is limited by the core diameter of the optical fibers used. The fiber-optic array detection scheme has been used to demonstrate the separation of dansylated amino acids (Fig. 6) [15]. A continuous separation of α-dansyl-l-arginine, Nε-dansyl-l-lysine, and dansyll-alanine is shown in Figure 6a. A single-diode electropherogram of the separation is shown in Figure 6b, indicating a well-resolved separation, with efficiencies ranging from 9500 to 26,000 theoretical plates [15]. Concentration and mass detection limits of 41 µM and 220 fmol, respectively, have been accomplished for α-dansyl-l-arginine with the fiber-optic detection method [27].

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Figure 6 (a) Continuous separation of three dansylated amino acids. A continuous injection of 0.5-mM samples of α-dansyl-l-arginine, Nε-dansyl-llysine, and dansyl-l-alanine in 25 mM CAPS buffer, pH 10.20, was made over 38 sec. The channel structure was 4.9 cm long with an internal height of 47.4 µm. The injection capillary was a 60-cm length of 144-µm-OD, 41-µm-ID capillary. The capillary and channel separation potentials were ⫹12 kV and ⫺550 V. The capillary was moved across the channel entrance for 350 steps @ 0.38 sec step. The channel separation current was 341 µA. (b) Electropherogram extracted from diode 250 from the data represented in Figure 6a. (From Ref. 28.)

Fluorescence Detection with a Charge Coupled Device An alternative fluorescence detection method for channel electrophoresis is the use of a charge-coupled device (CCD). While one early report employed an arc lamp to induce fluorescence [34], subsequent works have utilized a laser (Fig. 7). In this LIF scheme, light from a highpower argon ion laser is expanded, shaped into a line, and focused across the channel width. Fluorescence emission is collected at an angle normal to the excitation source and focused onto the CCD with a camera lens. Use of a CCD allows enhanced spatial resolution and the ability to detect analytes anywhere along the channel length.

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Figure 7 Laser-based fluorescence detection with a CCD camera for channel electrophoresis. (From Ref. 31.)

While fluorescence is a highly sensitive detection method, only a small percentage of compounds fluoresce natively. Thus, derivatization methods to expand the range of molecules that can be studied with this method are of interest. Most derivatization has been accomplished either prechannel [35,36] or in the channel itself [37]. One problem with such derivatization schemes has been that analytes can become multiply labeled, therefore altering migration rates and hence elution times. Figure 8 (a) Depiction of channel construction for postchannel derivatization. Two top plates are beveled at a 45° angle to create a well at the gap in which derivatization reagent can be placed. The channel is formed by adhering the two top plates to a single bottom plate with polymeric spacers. (b) Continuous separation and detection of arginine, glycine, and transferrin injected onto the channel system simultaneously. This time vs. pixel No. graph shows these analytes continuously separated over a span of approximately 10 min. Arginine eluted first, followed by glycine and then transferrin. The capillary was moved at a constant speed of 300 msec/step. The gap was 20 µm wide. Voltage across the capillary was ⫹15 kV; across the channel was ⫹1.5 kV. (From Ref. 43.)

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Postchannel derivatization has alleviated this problem by labeling analytes after the majority of separation but before detection occurs. In this detection scheme, a gap is constructed in the top plate near the channel exit (Fig. 8a) [43]. The derivatizing reagent is added to the gap whence it can diffuse into the channel, reacting with analytes as they pass. Separations of arginine, glycine, and transferrin have been performed in this manner using naphthalene dicarboxaldehyde (NDA)/ 2-mercapoethanol to derivatize the analytes (Fig. 8b) [43]. Postchannel derivatization not only prevents multiple labeling, but it also decreases the amount of derivatizing agent needed since the reagent is not added to the entire run buffer. Electrochemical Array Detection To expand the applicability of channel electrophoresis beyond natively fluorescent molecules and ones that can be easily derivatized, an electrochemical detection method has been developed. This approach is especially applicable to biological systems in which many compounds, such as catecholamine neurotransmitters, are electroactive. Other advantages of an electrochemical detection format include selectivity and sensitivity levels comparable to those of fluorescence [38]. To maintain spatial integrity of detection, an array of 100 platinum electrodes has been fabricated onto the bottom channel plate at the channel exit (Fig. 9) [39]. The array is constructed using standard

Figure 9 Schematic of the electrochemical array detector after a channel has been constructed. Although only five electrodes are depicted, the array actually consists of 100 individually addressed platinum electrodes positioned across the channel exit.

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lithographic techniques and involves depositing approximately 100 nm of platinum over a thin adhesion layer of titanium. Each resulting electrode is 1.5–2.0 mm long, 95-µm wide, and is separated from adjacent electrodes by 5 µm. The electrodes are held at a constant oxidation potential versus a reference electrode, while an auxiliary electrode is positioned in the exit reservoir to conduct current. As electroactive species migrate out of the channel and over an electrode, oxidation of the compounds occurs. The current from each electrode is amplified and subsequently collected by an A/D converter interfaced with a computer. The electrodes are isolated from the separation potential using the optimized end-column detection method described for CZE [44]. 6.3 CAPILLARY SAMPLE INTRODUCTION FOR DYNAMIC SEPARATIONS The coupling of channel electrophoresis with a capillary sample introduction format has been especially advantageous for examining dynamic systems such as cellular microenvironments and small reaction chambers. Since the size of the environment probed is limited only by the size of the sampling capillary, extremely small environments (as small as single nanoliters) can be examined. Moving the transfer end of the capillary back and forth across the channel entrance enables continuous monitoring of the sampling region for as long as desired. For example, the continuous sampling and separation of a solution of dopamine and catechol for nearly 40 min is shown in Figure 10 [16]. The amount of dopamine detected for each pass of the capillary over a single electrode is only 720 fmol injected in that width of channel, demonstrating the ability of this technique to perform sensitive, small-volume separations over extensive periods of time. 6.3.1

Channel Separation Techniques

A variety of techniques have been developed for channel electrophoresis that can be applied to the monitoring of dynamic systems. These include flow injection analysis, the use of ultranarrow channels, and the combinations of differentially selective separation modes.

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Figure 10 Continuous separation of 1.3 mM dopamine and 2.7 mM catechol with electrochemical array detection in an 8-µm-high channel structure of 4.8-cm length. The capillary ID was 15 µm. Field strengths were 310 V/ cm in the channel and 360 V/cm in the capillary. (From Ref. 16.)

Flow Injection Analysis (FIA) The ability of channel electrophoresis to probe dynamic environments has been demonstrated by continuously monitoring analytes in a flow injection system [28]. This technique, coupled with the fiber-optic array detection scheme described earlier, involves inserting the sampling capillary into a piece of 0.7-mm internal diameter (ID) medical grade

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Figure 11 Schematic of the capillary-channel system coupled to a flowing tube to demonstrate dynamic analysis. (From Ref. 28.)

tubing (Fig. 11). Injection is accomplished by applying electrophoresis potential to the tubing just downstream from the capillary. As analytes are swept down the flow tube, a small portion are injected into the capillary and migrate into the channel. When examining dynamic systems, it is important to be able to obtain qualitative and quantitative information about the analyte(s), as well as temporal characteristics of the region being probed (such as the duration of analyte appearance and concentration changes of analyte over time). These aspects have all been demonstrated with the FIA system. A sample plug consisting of four dansylated amino acids has been injected into the FIA apparatus while the capillary has continuously deposited material onto the separation channel. The resulting separation is plotted three-dimensionally to show the temporal resolution obtained (Fig. 12a). It is evident that aspartic acid has actually separated from the other compounds in the capillary, but the other three components have required the complete system to be resolved. In another experiment, intended to simulate the monitoring of changing analyte concentrations, sequential plugs with increasing arginine concentrations have been inserted into the flow injection stream [28]. Four regions of increasing intensity are observed in the resulting data, corresponding to the increasing concentrations in the FIA stream (Fig. 12b). Together, these experiments confirm the ability of channel electrophoresis to qualitatively and quantitatively monitor dynamic environments.

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Figure 12 Continuous separation of an analyte mixture sampled from a dynamic environment. (a) A 3-mL mixture of 3.00 mM α-dansyl-l-arginine (Arg), 3.06 mM Nε-dansyl-l-lysine (Lys), 3.20 mM dansylglycine (Gly), and 4.02 mM dansyl-l-aspartic acid (Asp) in 25.3 mM CAPS buffer (pH 9.94) was added to the flow injection reservoir to provide a 32-sec duration plug of analyte. Conditions: channel separation, ⫺1000 V and 0.84 mA; Capillary, ⫹12 kV and 57 cm length; capillary movement, 0.45 sec/25-µm step for a total of 450 steps across the channel and 450 steps back in the opposite direction. ‘‘I’’ represents an impurity. The capillary-to-channel linear flow velocity ratio was between 3 and 4. A three dimensional representation of the separation is shown. (b) Continuous electrophoresis of sequential additions of αdansyl-l-arginine to the flow injection reservoir. Analyte was added in four successive 4-mL plugs of 0.38-mM, 0.76-mM, 1.52-mM, and 3.04-mM analyte. The resulting three-dimensional plot is shown. (From Ref. 28.)

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Separations in Nanometer Structures The vast majority of work with channel electrophoresis has incorporated channels ranging in internal height from 8 to 109 µm. Miniaturization of the internal height is important for several reasons. First, the use of smaller heights permits the application of higher potential fields due to improved heat dissipation characteristics. Also, it is possible to obtain lower detection limits when performing separations in smaller channels [39]. Minimizing the channel internal height should improve the coulometric efficiency (mass detected/mass injected) since the channel exit is closer to the surface of the electrode array. Finally, coupling channels with small heights to similarly sized capillaries will permit the probing of even smaller dynamic microenvironments, since the capillary diameter defines the localized region that is sampled. Channels with heights as small as 8 µm have been constructed routinely using uniform glass microspheres. Smaller microspheres, however, have proven to be less uniform, resulting in larger channels than anticipated [40]. As an alternative, chromatographic stationary support particles have been applied as channel spacers due to their smaller variations in size. Submicrometer spacers have permitted the fabrication of 600 ⫾ 100 nm internal height channels as evaluated by scanning electron microscopy. The first continuous separation in a submicrometer channel has been demonstrated [40]. It is evident, especially when the capillary is slowly moved across the channel entrance, that sample overloading can prevent analyte resolution. This is largely a result of using a 15-µm capillary to transfer sample into the much smaller channel. Initial work coupling smaller capillaries (2–5 µm ID) to these channels has shown indications of improved resolving power (Fig. 13). Micellar Electrokinetic Channel Chromatography It has not always been possible to resolve complex mixtures completely using free-solution channel electrophoresis, especially when several of the compounds have similar structures. The addition of a surfactant to the channel running buffer leads to the formation of micelles, introducing a new mode of selectivity [45]. While electrophoresis still occurs in the channel, it is coupled with the varied partitioning of molecules into the micellar structures. This method, termed micellar electrokinetic

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Figure 13 Continuous separation of dopamine and catechol performed in a submicrometer-internal-height channel. The fused silica capillary (5-µm ID, 78.4-cm length) was moved in typewriter mode, with a rate of 0.4 sec/step in one direction and 0.15 sec/step on the return trip, with an applied potential of 25 kV. The capillary was filled first with a solution of 15.6 mM dopamine and 12.5 mM catechol, and was then switched to a solution of 25.0 mM dopamine and 20.0 mM catechol. The separation buffer was MES (25 mM, pH 6), with an applied channel voltage of 1.5 kV. Detection was at ⫹0.65 V vs. Ag/ AgCl. Material was deposited on the channel during 12 successive passes across the entrance. The increased intensity after the initial two passes results from the changed analyte concentrations being injected.

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channel chromatography (MEChC), has been combined with electrochemical array detection to separate five neurotransmitters not resolved via free-solution channel electrophoresis [19,41]. A continuous separation of a 12-min injection of catechol, norepinephrine, epinephrine, dopamine, and dihydroxybenzylamine in an 8-µm-high channel is shown in Figure 14. The ability to resolve these similar biological compounds demonstrates the promise of MEChC for investigating biological microenvironments. 2D Separations in Channels A multidimensional separation technique is often needed to obtain adequate resolution when analyzing complex samples. This requirement can be due to the limited peak capacity of a single dimension or simply because the separation mode is unable to separate certain analytes due to their physical similarities. Channel electrophoresis lends itself to multiple dimensions since the sample introduction capillary and channel can separate analytes using orthogonal separation modes. Replacing the CZE capillary with a packed micro-LC column for sample introduction into an electrophoresis channel allows separations to based sequentially on hydrophobicity and size/charge [46]. Liu and Sweedler [36] have similarly adapted the channel electrophoresis technology by coupling a CZE capillary (size/charge) with a polyacrylamide gel–filled channel (size). This technique has been used to provide two-dimensional separations of a peptide mixture, a tryptic digest of trypsinogen, and a portion of an individual Aplysia californica neuron. While not completely orthogonal separation modes, the combination of CZE with gel electrophoresis resolves components not separated with either technique individually. 6.3.2

Applications

To date, channel electrophoresis with capillary sample introduction has been utilized in a variety of dynamic sampling applications. The ability to perform parallel separations with continuous, low-volume sampling makes this technology ideal for performing kinetic assays or monitoring biological systems at a single-cell level.

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Kinetic Assays Channel electrophoresis, because of its ability to monitor multiple components continuously over long periods of time, is well suited to determination of the kinetic parameters of a dynamic reaction system. Liu and Sweedler [34,35] have demonstrated the applicability of this technique to kinetic assays. In one such experiment, a 200-nL solution of arginine, tryptophan, and NDA/CN⫺ (a derivatizing agent) has been injected onto the sampling capillary. The changing chemical composition of this solution as derivatization of both amino acids occurs has been examined continuously for approximately 4 min using the CCD detection scheme described previously (Fig. 15). These results clearly indicate the faster reaction kinetics for arginine derivatization versus the tryptophan derivatization, with a calculated first-order rate constant of 0.042 sec⫺1 [34]. A more complicated reaction system has also been examined, monitoring the enzyme catalyzed hydrolysis of fluorescein di-β-D-galactopyranoside (FDG) [35]. This hydrolysis leads to sequential formation of the fluorescent products fluorescein mono-β-galactoside (FMG) and fluorescein. Because of the continuous separations achieved by channel electrophoresis, the changing concentrations of both fluorescent products can be monitored (Fig. 16). These experiments demonstrate the ability of channel electrophoresis to quantitatively determine kinetic information in a dynamic system. Monitoring Release from Cells One of the most exciting applications of channel electrophoresis is the investigation of microenvironments in and around single cells. Previous work designed to monitor transmitter release from cells has been accomplished with a variety of electrochemical [47–49] and optical imaging [50,51] techniques. These methods have high temporal resolution, but frequently lack the ability to distinguish between multiple

Figure 14 Continuous micellar electrokinetic channel separation. Conditions: 10 mM sodium dodecylsulfate in buffer reservoirs; channel 8-µm internal height, 4.9 cm, 1500 V; capillary 10-µm ID, 62.5 cm, 28 kV. Elution order: catechol, norepinephrine, epinephrine, dihydroxybenzylamine, and dopamine. (From Ref. 19.)

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Figure 15 (a) Two-dimensional electropherogram of a ⬃200-nL solution of arginine (Arg), tryptophan (Trp), and NDA. Conditions: 25 mM CAPS, pH 10.20. A separation potential of ⫺1000 V was used. The channel current was 610 µA. (b) Maximum peak height of the Arg-CBI and Trp-CBI derivatives at each time in (a). (From Ref. 34.)

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Figure 16 Maximum peak height versus time for fluorescein mono-β-galactoside (FMG) and fluorescein obtained from a FDG hydrolysis solution (data for fluorescein are divided by 10 before display). The experimental conditions are as in Figure 15. (From Ref. 35.)

analytes. Microcolumn separation techniques have also been applied to cellular investigations [52,53]. The gains in chemical selectivity realized by these separation techniques, however, are usually offset by the loss of temporal information, thus limiting the ability to study individual release events which occur on a second to subsecond time scale. With the ability to perform extremely fast separations on chips [14], the possibility of gaining temporal information by performing frequent serial separations exists. However, sample introduction is still problematic with these systems. Channel electrophoresis with capillary sample introduction, with its combination of high temporal resolution and chemical selectivity, is ideal for this type of dynamic application. Preliminary investigations of release from bovine adrenal cells have been conducted [41]. In these experiments, one end of the sam-

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pling capillary is positioned directly next to a cluster of adrenal cells. A small volume of nicotine or potassium, known to stimulate epinephrine and norepinephrine exocytotic release from these cells, is added to the region immediately surrounding the cells and capillary tip. The capillary potential is then applied, causing solution outside the cells to be injected into the capillary and eventually into the separation channel. A long-duration signal indicative of neurotransmitter release is shown in Figure 17. The variation in peak intensity demonstrates the ability to monitor changing concentrations in biological microenvironments. The peak is presumed to be a combination of catecholamines, and refinements are under way to improve sensitivity and resolution. Liu and Sweedler [37] have introduced an adaptation to the sampling system for cellular studies. Instead of placing the capillary end near the cell(s) of interest, the cell is inserted into a nanoperfusion chamber constructed in the capillary itself. Pharmacological manipulation of the cell is easily achieved by flowing a small plug of the drug past the cell. Collection efficiency is increased greatly, since all released analyte is contained within the capillary. In addition, placing the perfusion chamber near the channel-injection end of the capillary improves time resolution by preventing significant diffusion or separation within the capillary. A perfusion chamber study, combined with a prechannel fluorescence derivatization scheme, has been used to monitor release from a cluster of isolated neurons from the cerebral ganglion of the salt water snail Aplysia californica [37]. A short plug of elevated KCl, a known exocytotic stimulant for Aplysia, has been passed over the cells while sample is deposited continuously onto the channel. Using this approach, two compounds with different electrophoretic mobilities have been observed. One of these analytes appears to be released for nearly 5 min after stimulation, while the other is released on a much shorter time scale (Fig. 18a). Exocytotic release has been confirmed by preceding the KCl plug with an injection of MgCl2, which blocks the Ca2⫹ channels and therefore inhibits neurotransmitter release. The chemical nature of the two resolved compounds remains unclear. Experiments using this method have also been performed to monitor the release from single Aplysia neurons [37]. Separation of a continuous 5-min injection yields three components (Fig. 18b). The authors

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Figure 17 Two-dimensional plot of an electropherogram obtained after stimulating a cluster of adrenal cells for 5 min with 100 mM potassium. The separation capillary was ⬃78-cm long and an injection voltage of 10 kV was applied throughout the duration of the experiment. The capillary was moved at a rate of 0.5 sec/step for 400 steps per pass across the entrance. The channel voltage was 500 V, resulting in a current of ⬃0.11 mA. The separation buffer in the channel reservoirs was ⬃10 mM TES, pH 5.7, with 10 mM SDS.

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Figure 18 (a) Electropherogram showing release from a group of cerebral ganglion neurons. The cells were positioned against a frit near the capillary exit, and a plug of KCl was passed over the cells to stimulate exocytosis. (b) An electropherogram from stimulating a single cerebral ganglion cell with elevated KCl in seawater. An Arg and Asp solution is injected 60 sec after the KCl injection as internal standards. (From Ref. 37.)

speculate that one compound, released almost immediately after stimulation for a duration of ⬃10 sec, is likely aspartate, based primarily on its similar electrophoretic mobility to a standard. The identities of the other components are unclear. These experiments represent an important first step in using channel technology to look at release from single cells. 6.4

CAPILLARY SAMPLE INTRODUCTION FOR PLUG/PARALLEL DNA SEPARATIONS

Channel electrophoresis with capillary sample introduction has also found application in the field of high-speed, low-volume DNA separa-

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tions. A wide variety of techniques have been used for such separations, including conventional slab gels, gel-filled capillaries [7], gel-filled capillary arrays [54], ultrathin slab gels [55], and sieving matrix separations. The use of microfabricated chips for the separation of DNA restriction fragments and sequencing reaction products has also received a great deal of recent attention [56,57]. Despite their widespread use, most of these methods are limited by difficulties in sample introduction. Channel electrophoresis is advantageous in that it combines the parallel processing of ultrathin slab gels with the ease of capillary sample introduction. This coupling makes possible a high throughput comparable to that of capillary array systems without the need for addressing large arrays of capillary entrances. The use of a single narrow-bore capillary also prevents gel overloading, thus allowing even narrower gels to be used with improved heat dissipation. In addition, this sample introduction system can be easily automated by moving the capillary into an array of sample chambers. As with the dynamic separations, sample is injected into and migrates through the buffer-filled capillary. For DNA it is assumed that no separation occurs in the capillary, since all of the species should have a similar mass-to-charge ratio regardless of size. Thus, an injection of DNA fragments with wide size variations should migrate through the capillary in a single plug. Translating the capillary across the channel entrance allows the deposition of multiple sample plugs onto the channel for separation. For detection, ethidium bromide is added to the run buffer. The intercalation of ethidium bromide with DNA as it migrates through the channel yields a large fluorescent signal that can be collected with a charge-coupled device as described previously. 6.4.1

Ultrathin Slab Gels

The channel structures used for ultrathin slab gels are similar to those constructed for dynamic separations described earlier. The channels are fabricated from quartz plates (16 cm long ⫻ 5 cm wide). Scotch tape along the lengths of the quartz plates is used as a spacer, forming a rectangular channel with an internal height of 57 µm. Prior to casting the slab gel, the quartz plates are soaked in a silane solution to ensure

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that the polyacrylamide will crosslink to the quartz plates and not migrate out of the channel. After rinsing the plates, a solution of acrylamide with polymerization initiators is poured onto one quartz plate with the tape spacers along the side. The second plate is immediately positioned on top of the first, allowing any excess solution to spill out of the channel. The plates are then bound together with epoxy along the edges while the gel polymerizes. Although methods to fill the channel with gel after channel construction have been developed, these techniques generally result in multiple bubbles, premature crosslinking, or difficulties in taking the channel apart. 6.4.2

Separations of DNA Restriction Fragments

Channel electrophoresis with an ultrathin slab gel has been used to separate restriction fragments from a variety of double-stranded DNA samples [31–33]. Multiple injections of the same fragment set have been made on the slab gel with high resolution and no indications of sample carry over from plug to plug [31]. A separation of 12 discrete injections of HaeIII-digested ΦX174, PBR322, and PUC18, each deposited four times onto a slab gel, is shown in Figure 19a. These digests range in size from 8 to 1353 base pairs. Each injection into the capillary is made at intervals of 3 min, and the space between lanes is defined by the buffer spacing between injections. As with the dynamic separations discussed earlier, the diagonal nature of the separation lanes results from the uncorrected time bias of sample introduction. Correction is easily accomplished by subtracting the time difference. A standard electropherogram obtained at a single pixel shows the resolution obtained in the separation of the digested ΦX174 (Fig. 19b).

Figure 19 (a) Twelve plug separations of HaeIII-digested ΦX174, PBR322, and PUC18 DNA. The 6% polyacrylamide gel was 57 µm thick, with a total gel length of 16 cm (11.5 cm to the detector). The separation potential was 125 V/cm. The capillary was 50.3 cm long, with a tip of 90 µm; Vcap ⫽ 15 kV. Injections: 3 sec at 15 kV every 3 min. (b) Signal versus time response at a single pixel from data similar to that shown in (a). The data shown correspond to fragments of HaeIII-digested ΦX174 DNA. (From Ref. 31.)

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The time axis represents the time needed for fragments to travel through the entire separation system, including the capillary. The channel separation itself occurs in 20–30 min at the potentials applied. Nine of the 11 fragments of ΦX174 are baseline resolved, and the 10-base difference between fragments 271 and 281 is clearly distinguishable. The resolution obtained in the separation is typical for the level expected for dsDNA in a 6% gel [31]. The throughput of channel separations is dependent primarily on the lateral resolution (i.e., across the width of the channel) of each injection. With narrower injection plugs, a larger number of samples can be deposited across the channel entrance. For example, if lateral dispersion can be limited to 0.18 mm, then ⬎200 DNA separations could be accomplished simultaneously in a 4-cm-wide channel [16]. To date, most sample plugs have resulted in a 1-mm plug width near the channel exit [32]. By focusing the detector optics at the entrance to the gel, it has been determined that the injection plug itself is also about 1 mm in width, indicating that little lateral broadening is occurring within the gel. These results suggest that the broadening must be occurring at the interface between the capillary (50 µm ID) and the gel. Depending on the exact geometry of the beveled entrance to the channel, the position of the capillary, and the size of the capillary tip (which can be modified by HF etching), the gap between the capillary and channel can be ⬎20 µm [19]. Maintaining a lower potential field in the capillary versus in the channel minimizes band broadening in this gap. In addition, the position and structural integrity of the capillary tip are very important in minimizing injection widths [31]. In fact, experiments have indicated that with a cracked capillary tip, a substantial portion of eluting sample does not even enter the channel. Alternative means of controlling lateral dispersion are being explored. One of the primary advantages of utilizing capillary sample introduction is its low-volume sampling capability. The ability to isolate DNA into small vials, perform reactions on the nanoliter scale, and analyze the results has significant implications for future genomic research. The sampling limits of the channel injection scheme have been tested by injecting HaeIII-digested ΦX174 DNA from 1-nL vials [33]. These nanovials have been constructed on silicon wafers using standard lithographic and etching procedures, resulting in arrays of square py-

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ramidal containers. Injection has been accomplished electrokinetically by first coating the silicon with a layer of gold or platinum and then applying a potential to the wafer. To prevent sample evaporation, which can occur in just seconds with such small volumes, a small amount of glycerol has been injected into the vials with the DNA samples. Baseline resolution of 10 fragments has been obtained for the ΦX174 DNA digest when sampling from these vials, demonstrating the applicability of channel electrophoresis to DNA analysis from nanoliter volumes.

Figure 20 Nine sequential separations of 3 ladders and six different STR samples. A 6% polyacrylamide gel, 57 µm thick, was used with a detection window at 14 cm. A 46-cm-long, 50-µm-ID transfer capillary was used at 10 kV (218 V/cm) following hydrodynamic injection (approximately 9 nL). The separation field in the ultrathin slab gel was 100 V/cm. Ethidium bromide was used only in the separation buffer at a concentration of 1.3 µM. From the right side of the figure: lane (1) ladder; (2) alleles 14 and 17; (3) allele 17, 17; (4) alleles 15 and 16; (5) ladder; (6) allele 17, 18; (7) allele 17, 17; (8) alleles 16 and 17; (9) ladder. (From Ref. 30.)

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Channel Separations for Analysis of Short Tandem Repeats

Channel electrophoresis in ultrathin slab gels has been applied to the analysis of polymerase chain reaction amplified short tandem repeat (STR) samples [30]. STRs consist of repeated units of DNA ranging in size from 3 to 7 bp. They are characterized by high variation in the number of repeating units and as such are increasingly used as genetic markers. Nine sequential separations of three DNA ladders and six different STR samples across 1.5 cm of an ultrathin slab gel are shown in Figure 20. Although the resolution is not yet optimized to resolve STRs containing four-base repeats, coupling capillary sample introduction to slab gels should allow for higher potentials and faster separations than more standard methods while maintaining high throughput.

6.5

FUTURE DIRECTIONS

Channel electrophoresis with capillary sample introduction has already demonstrated applicability to a wide variety of analyses. In the area of dynamic separations, the technique has been applied to the monitoring of a flow injection analysis stream and the examination of reaction kinetics. The potential to study release from biological systems at a single-cell level has also been examined. Through the application of more selective separation modes such as MEChC or multidimensional techniques involving multiple separation modes, resolution improvements are readily obtainable. These techniques should enhance the abilities to distinguish and identify compounds contained in and released from single cells. In addition, the use of smaller capillaries for sample introduction will not only lead to better sample transfer to submicron channels but will also enable smaller microenvironments to be probed. With a small enough capillary, it would even be possible to continuously sample from within the cytoplasm of a single cell, making possible the investigation of dynamic pharmacological effects on the cellular interior. The combination of low-volume sampling, fast separation, and high throughput has also made channel electrophoresis very amenable to DNA analysis. Although ultrathin slab gels have comprised most of

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the work in this area, some efforts using the sieving buffer hydroxy ethyl cellulose have been reported [29]. The use of polymeric sieving matrices instead of gels eases channel filling and extends the lifetime of the channel. Optimization of both double-stranded and single-stranded DNA fragment separations are under way. In addition, efforts to limit lateral diffusion in the capillary/channel interface are in progress for all types of channel separations. ACKNOWLEDGMENTS This work was funded, in part, by grants from the National Science Foundation and the National Institutes of Health. We would like to acknowledge all our past and present coworkers whose work is cited in this chapter. We also thank Don Cannon, Tom Colliver, and Julie Lapos for helpful comments and discussion on this manuscript. Steven W. Suljak was the recipient of a National Science Foundation Graduate Research Fellowship. REFERENCES 1. Beale, S. C. Anal. Chem. 1998, 70, 279R–300R. 2. Kennedy, R. T., Oates, M. D., Cooper, B. R., Nickerson, B., Jorgenson, J. W. Science 1989, 246, 58–63. 3. Ewing, A. G. J. Neurosci. Methods 1993, 48, 215–224. 4. Lee, T. T., Yeung, E. S. Anal. Chem. 1992, 64, 3045–3051. 5. Jankowski, J. A., Tracht, S., Sweedler, J. V. Trends. Anal. Chem. 1995, 14, 170–176. 6. Gilman, S. D., Ewing, A. G. J. Cap. Elec. 1995, 2, 1–13. 7. Guttman, A., Cohen, A. S., Heiger, D. N., Karger, B. L. Anal. Chem. 1990, 62, 137–141. 8. Manz, A., Harrison, D. J., Verpoorte, E. M. J., Fettinger, J. C., Paulus, A., Ludi, H., Widmer, H. M. J. Chromatogr. 1992, 593, 253–258. 9. Effenhauser, C. S., Bruin, G. J. M., Paulus, A. Electrophoresis 1997, 18, 2203–2213. 10. Colyer, C. L., Tang, T., Chiem, N., Harrison, D. J. Electrophoresis 1997, 18, 1733–1741. 11. Jacobson, S. C., Hergenroder, R., Moore, A. W., Ramsey, J. M. Anal. Chem. 1994, 66, 4127–4132.

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7 Electric-Field-Mediated Separation of DNA Fragments on Planar Microgels Andra´s Guttman Torrey Mesa Research Institute, San Diego, California

7.1 INTRODUCTION Throughput is one of the greatest single factors impacting the costeffectiveness of large-scale genetic mapping, mutation (SNP) detection, or PCR product analysis by using conventional agarose or polyacrylamide gel electrophoresis systems [1]. Despite numerous refinements in electrophoresis techniques over the past decade, the process is still not efficient enough and hardly automated. The time required to preparing, loading, and separating DNA fragments using conventional gel electrophoresis, and staining and exposure times are all added up. Although several attempts have been made to automate this technology [2], DNA fragment analysis in most laboratories is still done in a very conventional way: using slab gel electrophoresis separation with ethidium bromide staining [3], or more recently with novel, higher-sensitivity fluorophores [4]. The introduction of fluorescent dyes provided the possibility of employing regular or electronic (CCD) cameras to take pictures of transilluminated gels for data evaluation and archiving [5]. By all means, the existing technology of separating DNA 165

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fragments using gel electrophoresis is a task that requires multiple labor-intensive steps, such as gel casting, sample loading, staining and imaging/documentation/data evaluation. These tasks are not readily integrated and automated for high-throughput applications. Some of the large, automated DNA sequencing systems have recently been reported to accommodate genotyping and STR profiling [6] using crosslinked polyacrylamide gels and fluorescently labeled primers; however, the configuration of these devices does not accommodate the use of agarose gels as separation medium. In addition, all the reported methods on those commercially available automated electrophoresis systems require preseparation covalent fluorophore labeling. Electrophoretic separation of DNA fragments, such as PCR products, is usually accomplished in agarose, polyacrylamide, or composite agarose-polyacrylamide gels [7]. At slightly basic separation conditions (7 ⬍ pH ⬍ 9), DNA molecules are negatively charged, so they should be loaded at the cathode end of the separation platform and migrate toward the anode when the electric field is applied. Under denaturing conditions the electrophoretic mobility of DNA fragments is primarily determined by their size, while under nondenaturing separation conditions it is also influenced by the sequence-dependent secondary structure [8]. Agarose gels are usually employed to analyze double-stranded DNA molecules in size from hundreds of base pairs (bp) to tens of thousands of base pairs, and polyacrylamide gels are regularly used for high-resolution DNA fragment analysis from several base pairs up to 1000 bp. Attempts to discover faster and higher-resolution electrophoresis separation techniques and mediums started as early as in the 1950s with system miniaturization [9]. Edstrom first described microelectrophoresis separation of picogram scale nucleic acid bases from single cells along a silk thread [10]. A decade later, Matioli and Niewisch [11] developed a microelectrophoresis method for the separation of hemoglobins from individual erythrocytes using polyacrylamide fibers. They attained very rapid separations by applying electric fields as high as 1000 V/cm. The reason these microelectrophoretic methods were not exploited to a greater extent after the initial demonstrations was probably due to the inadequacy of the imaging techniques at that time, i.e., to capture separations in such a minute scale. Later, Neuhoff [12]

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employed microelectrophoretic methods using 2-mm-long polyacrylamide gels in narrow-bore glass tubes to separate both DNA and RNA molecules. His progress was ultimately hampered by the inability to control extra Joule heat during the application of the extremely high separation voltages. The relationships between gel dimensions and the accumulations of Joule heat are now fully understood. First Karger and coworkers [13] and later Grossmann et al. [14] showed that one of the key factors influencing the extent of Joule heat is the thickness of the separation platform. Electrophoresis in capillary dimensions proved to be much less susceptible to effects of Joule heat, because of the ability to dissipate heat via the large surface areas typical of capillary columns. Obtaining high surface-to-volume ratios in planar format electrophoretic systems requires very thin layers, preferably reaching the thickness of capillary dimension. Separation platforms no thicker than 0.25 mm allow the application of high electric field strengths (typically ⬎ 50 V/cm), enabling rapid and efficient electrophoretic separations of DNA fragments [15]. Note that in many current applications, including most DNA fragment analyses, the necessary decrease in sample volume required by the planar microchip format has limited its use, due to the lack of appropriate injection methodology. However, recently introduced membranemediated loading technology successfully addressed this issue [16]. Recently emerging microfluidics-based analytical techniques [17] brought the promise to further the speed and throughput of electricfield-mediated separations. Microfluidics devices enable the so-called cross-channel injection which is a precisely controlled injection form to load picoliter amounts for short separation distances with the capability of multiplexing [18]. Conventional gel electrophoresis separations of DNA fragments rely on complex relationship among several factors including gel pore size, electric field strength, ionic moiety, buffer composition, and conductivity, as well as the type of the migration of DNA fragments through the gel matrix. Smaller DNA fragments migrate by tumbling through the pores with much larger average radii than that of the radius of gyration of the fragments; thus, they become size separated on a basis of time required to find their path through the pores of the gel matrix [19]. Larger fragments, i.e., whose radii of gyration are larger

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that of the average pore size of the gel, become elongated toward the electric field in order to migrate through the smaller pores. This phenomenon is called reptation [20], which is mainly induced through increases in either the gel concentration (i.e., decreasing the pore size) or the applied electric field strength, and in extreme instances may even result in size independent migration of the analyte molecules. The most recent advances in the electrophoretic separation of nucleic acids come from exploration of new separation matrices. Linear polymers, such as noncrosslinked polyacrylamide [21], derivatized celluloses [22], and polyethylene oxides [23], have all been widely demonstrated to be effective in electric-field-mediated size separation of biopolymers. The advantages of these noncrosslinked polymers have been demonstrated almost entirely in high-performance capillary electrophoresis applications [24–26], albeit it has been shown that very high-concentration linear polymers can be used in planar separation format [27]. Agarose gel–filled capillaries were extensively studied by Bocek and Chrambach [28] for the separation of dsDNA molecules. Chemically modified agarose gels or composite agarose—noncrosslinked polymer gels capable of resolving several base pair differences in DNA fragments of several hundreds of base pairs in length–have also been developed [29]. The employment of noncrosslinked, linear polymers for DNA fragment analysis applications may be advantageous in several respects. First, it has been shown that noncrosslinked polymers may be supplied in a desiccated, dry form, providing long shelf life [30]. Second, planar form linear polymers can be rehydrated to any of the range of final gel concentrations, buffer compositions, and/or ionic strengths [31]. Note that lower-viscosity noncrosslinked polymers can be easily replaced within the separation platform, so planar microchips supporting repetitive work can be readily used with these matrices [32]. Visualization of nucleic acids by covalent labeling with fluorescent tags, such as fluorescein, tetramethyl-rhodamine, Texas Red etc., has been practiced for years [33]. This approach can be used for highsensitivity DNA fragment analysis provided the analyte is covalently labeled by the appropriate fluorophore prior to the separation process. The approach reported here employs noncovalent affinity binding (e.g., intercalation) dyes for in migratio labeling of the dsDNA fragments

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during their electrophoretic separation. This method is beneficial in two ways. First, unlabeled DNA fragments can be readily visualized since they become labeled during the separation process. Second, the complexation phenomenon usually increases the separation selectivity, resulting in higher resolution [34]. 7.2 SEPARATION PLATFORM Figure 1 shows the picture of the automated planar microgel electrophoresis system consisting of a high-voltage power supply (1), ultrathin-layer casette with built-in buffer reservoirs (2), and a fiber-optic bundle-based scanning detection system (3). A lens set (4) connected to the illumination/detection system via the fiber-optic bundle scans across the gel by means of a high-speed translation stage (5). A laser excitation source (6) and an avalanche photodiode (7) are connected to the central excitation fiber and the surrounding collecting fibers of the fiber-optic bundle, respectively [35]. Interface electronics (8) is used to digitize the analog output of the detector and to connect the system to a personal computer (9). The horizontal assembly includes a positional heat sink to hold the gel-filled casette (10) and also to eliminate local heat-spot-generated separation irregularities by means of homo-

Figure 1 Automated planar microgel electrophoresis system.

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geneous dissipation of any extra heat over the gel surface during the separation. Three heating cartridges and a high-speed fan control the temperature (above ambient) of the separation platform at a preprogrammed level. 7.3

OPTICAL PATHWAY

Figure 2 shows the detailed block diagram of the optical pathway of the illumination/detection system. The use of a relatively small numeric aperture illuminating fiber allows the light to be launched into a reasonably small conical angle [35]. A 532-nm frequency doubled Nd-YAG laser (1) is used as a light source which is electronically modulated at 5 kHz to facilitate lock-in detection of the signal. The beam is collimated by an aspheric lens (2) and the collimated light beam is sent through a narrow-band interference filter (3), which cleans up the beam

Figure 2 Optical pathway of the illumination/detection system. Inset: crosssectional view of the fiber-optics bundle.

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by blocking wavelengths which interfere with the emission light wavelengths. An aspheric lens (4) is used to focus this beam onto the face of the excitation fiber (5). The longer focal length of this lens, along with the small diameter of the input light beam, ensures that the light enters into the fiber with a small numerical aperture. The fiber-optic bundle (11) comprises an illumination fiber (5) and six collection fibers (6) which surround the illuminating fiber (see inset in Fig. 2). The scanning lens set has a pinhole filter between the collimating and objective lenses (13) providing further filtering of ambient light and laser scatter. The lens in front of the fiber-optic bundle is a spherical plano lens (12). A diffraction-limited aspherical lens (14) is used to focus the beam onto the separation gel (15). As the inset in Figure 2 depicts, in this particular arrangement the excitation fiber (5) is placed in the geometric center of the lens set (12–14), and the collecting fibers (6) are located off the geometric center of the lens set. To collect the fluorescent signal emitted by the sample, a large aspheric (7) lens is used which forms a collimated beam that is sent through an interference filter (8), passing only a 25-nm-wide band in the center of the emitted light spectra. A similar type aspheric lens (9) is used to focus the light beam onto the avalanche photodiode detector (10). 7.4 PLANAR MICROGELS The reuseable separation planar microgel is shown in Figure 3 having a flat bottom and top plates (1) joined and secured in parallel, spaced apart 190 µm with buffer reservoirs permanently fixed to both ends (2). Two spacers (3) run along the inner face edges of the plates to assure consistent distance between the glass plates. For filling the chip a syringe (6) is used with a sealing applicator nozzle (5) that matches to the top of the buffer reservoirs (4). To introduce the melted agarose, the nozzle is placed at the top of one of the reservoirs, and the gel is pumped into the platform. In order to prevent premature solidification of the separation matrix, preheating of the planar microgel to approximately 40–45°C prior to the introduction of the melted agarose gel is strongly suggested. Appropriate amount of DNA staining dye, e.g., ethidium bromide (10–50 nM), is added to the melted agarose solution just before filling into the planar microgel.

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Figure 3 Planar microgel and the refilling assembly.

7.5

MEMBRANE-MEDIATED SAMPLE LOADING

Membrane-mediated sample loading provides a reliable and easy loading mechanism for ultrathin planar microgel that is also applicable for both vertical and horizontal formats [36]. The samples are spotted directly to the surface of the loading membrane tabs, manually or automatically (robots), outside of the separation/detection platform. The sample-spotted membrane is then placed into the injection (cathode) side of the planar microgel, in intimate contact with the straight gel edge. By the application of the electric field, the sample components migrate into the gel. There is no need to form individual injection wells in the separation gel, and loading is accomplished easily on the bench top, outside of the separation platform. This novel sample injection method could also be readily applied to most high-throughput thinlayer slab gel electrophoresis–based DNA analysis applications (e.g., automated DNA sequencing [37]). 7.6

IN MIGRATIO FLUOROPHORE LABELING

Noncovalent, instantaneous, fluorophore labeling of DNA fragments by intercalation expands the detection sensitivity and separation potential of gel electrophoresis [38]. Complex formation with fluorogenic

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stains enables in migratio labeling, that is, the migrating DNA fragments (negatively charged) complex with the countermigrating fluorophore intercalator dye (positively charged) which is dissolved in the separation matrix. Using this method, high-sensitivity detection of the migrating DNA fragments, as well as high resolution of closely migrating fragments can be simultaneously obtained over a broad molecularweight range. The complexing dye (ligand: L⫹) intercalates between the strands of double-stranded DNA (P n⫺ ) molecules, and due to its positive charge (at the separation pH of 8.4), it results in decreased electrophoretic mobility of the DNA-ligand complex (PLm(n⫺m)⫺) by reducing its charge-to-mass ratio: P n⫺ ⫹ mL ⫹ s PLm(n⫺m)⫺ K⫽

(1)

(n⫺m)⫺

[PLm ] n⫺ ⫹ m [P ] ⋅ [L ]

(2)

where K is the formation constant of the complex, m is the number of the positively charged fluorescent dye molecules in the complex, and n is the total number of negative charges on the DNA molecules (in this instance, equal to the number of phosphate groups). Doublestranded DNA molecules have approximately one ethidium bromide binding site for five base pairs, slightly influenced by the dye/polymer ratio and salt concentration but there is no reported base composition selectivity of this binding [39]. Please note that while, for example, ethidium bromide affects the mobility and resolution of doublestranded DNA molecules, it has no significant effect on the migration and separation of single-stranded oligonucleotides. The appropriate concentration and type of the fluorophore complexing dye should be optimized for the excitation laser used in the illumination/detection system. For a 532-nm laser, ethidium bromide provides a good match (EX max ⫽ 512 nm), but other high-sensitivity fluorescent dyes can also be used. 7.7 AGAROSE-BASED REPLACEABLE SEPARATION MATRIX Agarose gels are characterized as large pore size, high mechanical strength, and biologically inert separation matrices [40]. As agarose is

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the medium of choice for the separation of relatively large dsDNA molecules with conventional slab gel electrophoresis, it can also be used to separate DNA in capillary dimensions as was reported earlier by Compton and Brownlee [41]. Since then, other research groups [42] have also reported promising results with agarose-filled capillaries using purified grades of low-electroendosmosis (EEO) agaroses to avoid electro-osmotic flow-mediated disruption of the separation in capillary dimensions. In the planar microgel format described in this chapter, dsDNA fragment analysis was accomplished using 2% agarose gel, filled into the separation platform at 60°C and used for electrophoresis separation after solidification at room temperature. One of the main advantages of employing liquefied agarose above its gelling temperature is that the gel can be easily replaced by simply pumping fresh melted gel into the platform; i.e., it can be easily filled, rinsed, and refilled. The inner surface of the planar microgel was coated with linear polyacrylamide in order to avoid the formation of an electric double layer and concomitant electro-osmotic flow generation [43]. Figure 4 exhibits the separation of various dsDNA ladder standards and restriction digest fragments over a broad size range of 50 bp to 23,000 bp, using a single agarose gel composition within the planar microgel. Panels A, B, and C depict rapid separations of the 50- and 100-bp and 200-DNA ladder standards in 25 min, respectively. Note that since the various ladder standards were generated by ligating sections of different sequences, some of the very same chain length fragments from the various ladders exhibited slightly different migration times, probably due to secondary structure differences recognized by the high resolving power of the separation system. Panel D shows a nice separation of the ϕX174 DNA Hae-III restriction digest mixture with particularly good separation between the landmark 271 and 281 bp fragments. Panel E depicts a rapid, high-resolution separation of the pBR322 DNA Msp-I restriction digest mixture. The high separation efficiency of the system enabled here to obtain even a challenging 4-bp resolution between the 238- and 242-bp fragments. Panel F exhibits the separation of the lambda DNA Hind-III restriction digest fragments ranging up to 23,130 bp in size. The largest peaks seem to be overloaded, probably due to the proportion-

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Figure 4 Separation of standard DNA ladders and restriction digest fragment mixtures by planar microgel-based agarose gel electrophoresis. (A) 50 bp, (B) 100 bp, and (C) 200 bp DNA sizing ladders. (D) ϕX174 DNA HaeIII, (E) pBR322 DNA Msp-I, and lambda DNA Hind-III restriction digest mixtures. Conditions: Effective separation length: 6 cm; separation matrix: 2% agarose gel in 0.5 ⫻ TBE buffer containing 25 nM ethidium bromide; running buffer: 0.5 ⫻ TBE; applied voltage: 750 V; temperature: 25°C; injection: membrane-mediated, 0.5-µL sample per tab.

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Figure 5 Detection linearity of the planar microgel system.

ally higher amount of ethidium bromide intercalated into those fragment. Quantitation and detection linearity was evaluated by injecting a series of dilutions of the 100-bp ladder from 25, 10, 5, 2.5, and 1.25 ng total DNA injected, corresponding to 0.863, 0.30, 0.17, 0.08, and 0.04 ng DNA per peak, respectively (Fig. 5). Based on these results, the limit of detection (LOD) of the automated planar microgel-based electrophoresis system was found to be ⬃0.08 ng DNA per band. Figure 6 Rapid genotyping of the DRD4 receptor gene by multiplexed (24) analysis of PCR products on planar microgels. Conditions: 2% agarose gel in 45 mM Tris, 45 mM boric acid, 1 mM EDTA, pH 8.4 (0.5 ⫻ TBE) buffer containing 50 nM ethidium bromide; running buffer: 0.5 ⫻ TBE; applied voltage: 750 V; effective separation length: 2 cm; current: 8 mA; temperature: ambient.

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Figure 6 depicts an example of rapid and simultaneous analysis of 24 PCR-generated genotyping samples in a highly multiplexed format on a planar microgel. It has been reported that such human behavioral traits as novelty seeking, hyperactivity disorders, and substance abuse may possibly be associated with the presence of a certain 48-bp variable number tandem repeat polymorphism of the human dopamine D4 receptor gene (DRD4). A highly optimized genotyping method was elaborated, allowing simultaneous PCR amplification with large differences in amplicon sizes (379 bp vs. 667 bp), substituting 50% dGTP by dITP and using nanograms of DNA template. High-throughput separation and instant visualization of the PCR products were accomplished within 10 min by automated planar microgel analysis. The electropherograms in Figure 6 show rapid detection of a wide range of possible genotypes (2x–8x). This includes such rear heterozygotes as the 2x and 8x 48-bp repeats in the same sample, proving the reliability of this novel technique that even enables detection of the longer alleles in the presence of shorter alleles [44]. The high sensitivity of the laser-induced fluorescence detection for the ethidium bromide–stained DNA fragments was especially important where deoxynucleotide analogs were used during PCR amplification. This method provides reliable and high-throughput genotyping of the variable number repeat region of DRD4 gene that leads to better understanding of genetic factors in normal and pathological human behavior. 7.8

CONCLUSION

A miniaturized DNA fragment analyzer has been introduced for highthroughput DNA fragment analysis using a multilane (virtual channels) configuration, employing replaceable agarose gels in conjunction with a reusable planar microgel format. Sample loading was accomplished by membrane-mediated loading technology, which also enabled robotic spotting of large number of samples [45]. The sample loading membranes were also bar-coded for easier identification and cataloging purposes, and could be stored for several days between spotting and analysis. The analyte DNA fragments were labeled in migratio (during the separation process) by complexation with fluorogenic intercalator or

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by conventional preseparation covalent labeling. The migrating bands were detected by the scanning fiber-optic bundle based integrated laserinduced fluorescence avalanche photodiode imaging system. Detection of the separated bands was accomplished in real time, by continuous scanning at just a few centimeters from the injection site on the multilane separation platform. It is important to note here that the detection of the migrating DNA fragments were accomplished in a timely basis, similar to capillary electrophoresis (CE) and high-performance liquid chromatography (HPLC), in contrast to the traditional spatial-based detection in agarose slab gel electrophoresis. This guaranteed that all DNA fragments in the sample mixture traveled the same distance from the injection point to the detection line. Cost-effectiveness for large-scale applications was assured by reTable 1 Comparison of Separation Performance and Sensitivity of the Planar Microgel Electrophoresis (Automated) and the Manual Submarine Slab Gel Electrophoresis Systems DNA fragment analysis Feature

Planar microgel

Manual

Gel preparation and loading Separation Staining/destaining Imaging/evaluation Total time

5 min

35 min

5 min NONE NONE 15 min

60 min 30 min 15 min 155 min

Number of lanes Time per sample

32 0.32 min

14 10 min

Sensitivity (ethidium bromide) Separation range (single gel) Required buffer volume Required gel volume

0.04 ng/band

0.2 ng/band*

20bp– 25,000bp 20 ml 1 ml

50–500, 500–5K, 5K–50K 250 ml 20 ml

* With UV transillumination and CCD camera detection imaging.

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using the same planar microgel or by simple replacement of the separation matrix by pumping fresh melted gel composition into the casette for consequent analyses. Table 1 compares the separation performance and detection sensitivity of conventional submarine agarose gel electrophoresis to the planar microgel electrophoresis platform, exhibiting significantly faster and higher-sensitivity analysis for the latter. Our goal is to develop an integrated microgel-based, automated, high-throughput DNA fragment analyzer by coupling sample preparation, PCR, restriction digestion, loading, and separation with data analysis and map construction. ACKNOWLEDGMENTS The author gratefully acknowledges the technical help of Loi Nguyen, Doug Evans, Swarna Ramanjulu, Nick Wilder, Greg Theriault, and Zsolt Ronai. REFERENCES 1. U.S. Department of Health and Human Services. The human genome project: new tools for tomorrow’s research. U.S. Gov. Print. Office 281– 846: 40022 (1991). 2. Gombocz, E., and Cortez, E. Appl. Theor. Electrophoresis, 4, 197 (1995). 3. Sambrook, J., Fritch, E.F., and Maniatis, T. Molecular Cloning, 2nd ed., Cold Spring Harbor Lab. Press, Plainview, NY (1987). 4. Haugland, R.P.H. In: Handbook of Fluorescent Probes and Research Chemicals, ed. M.T.Z. Spence. Mol. Probes, Inc., Eugene, OR, Chap. 8 (1996). 5. Southerland, J.C. In: Advances in Electrophoresis (eds: Chrambach, A, Dunn, M.J., Radola, B.J.) VCH Publishers, Weinheim, Germany (1993). 6. Ballard, L.W. http:/ /www.medstv.unimelb.edu.au/ABRFNews/1997/ September 1997. 7. Rickwood, D., and Hames, B.D. Gel Electrophoresis of Nucleic Acids, Oxford University Press, Oxford, England (1990). 8. Chrambach, A. Practice of Quantitative Gel Electrophoresis, VCH, Deerfield Beach, FL (1985). 9. Andrews, A.T. Electrophoresis, 2nd ed. Clarendon Press, Oxford, UK (1986).

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10. Edstrom, J.E. Biochim. Biophys. Acta 22, 378 (1956). 11. Matioli, G.T., and Niewisch, H.B. Science 150, 1824 (1965). 12. Neuhoff, V. Micromethods in Molecular Biology. Springer-Verlag, New York, (1973). 13. Nelson, R.J., Paulus, A., Cohen, A.S., Guttman, A., and Karger, B.L. J. Chromatogr. 480, 111 (1989). 14. Grossman, P.D., Menchen, S., and Hershery, D. Gene Anal. Tech. Appl. 9, 9 (1992). 15. Brumley, R.L., and Smith, L.M. Nucl. Acids Res. 19, 4121 (1991). 16. Guttman, A., Barta, CS., Szoke, M., Sasvari-Szekely, M., and Kalasz, H. J. Chromatogr. 828, 481 (1998). 17. Jacobson, S.C., Ramsey, J.M. In ‘‘High Performance Capillary Electrophoresis’’ (ed: M.G. Khaledi) John Wiley & Sons, New York, 1998. 18. Dolnik, V., Liu, S., Jovanovich, S. Electrophoresis, 21, 41, (2000). 19. Ogston, A.G. Trans. Faraday. Soc., 54, 1754 (1958). 20. Lumpkin, O.J., DeJardin, P., and Zimm, B.H. Biopolymers 24, 1573 (1985). 21. Heiger, D.N., Cohen, A.S., and Karger, B.L. J. Chromatogr. 516, 33 (1990). 22. Schwartz, H.E., Ulfelder, K. et al., J. Chromatogr. 559, 267 (1991). 23. Fung, E.N., and Yeung, E.S. Anal. Chem. 67, 1913 (1995). 24. Peacock, A.C., and Dingman, C.W. Biochemistry 7, 668 (1968). 25. Guttman, A. In: Handbook of Capillary Electrophoresis (ed. Landers, J.P.), CRC Press, Boca Raton, FL (1994). 26. Tietz, D., Gottlieb, M.H., Fawcett, J.S., and Chrambach, A. Electrophoresis 7, 217 (1986). 27. Bode, H.J. Electrophoresis ’79; Walter de Gruyter & Co., New York, 1980, p. 39. 28. Bocek, P., and Chrambach, A. Electrophoresis, 12, 1059 (1991). 29. Soto, D., and Sukumar, S. PCR Meth. Appl. 2, 96 (1992). 30. Schwatz, H. and Guttman, A. Separation of DNA by Capillary electrophoresis, Primer 5. Beckman Instruments, Inc., Fullerton, CA (1995). 31. Allen, C.R., Graves, G., and Brudowle, B. BioTechniques, 7, 736 (1989). 32. Guttman, A. Trends Anal. Chem. 18, 694, 1999. 33. Smith, L.M., Sanders, J.Z., Kaiser, R.J., Hughes, P., Dodd, C., Connell, C.R., Heiner, C., Kent, S.B.H., and Hood, L.E. Nature 321, 674 (1986). 34. Guttman, A., and Cooke, N. Anal. Chem., 63, 2038 (1991). 35. Trost, P., and Guttman, A. Anal. Chem., 70, 3930 (1998). 36. Cassel, S., and Guttman, A. Electrophoresis, 19, 1341 (1998).

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37. Gerstner, A., Sasvari-Szekely, M., Kalasz, H., Guttman, A. BioTechniques 28, 628 (2000). 38. Guttman, A. In: HPLC, Practical and Industrial Applications (ed. Swadesh, J.). CRC Press, Boca Raton, FL, (1997), p. 323. 39. LePecq, J.B., and Paleotti, C. Anal. Biochem., 17, 100 (1966). 40. FMC. MetaPhor Agarose. FMC Bioproducts Application Note (1992). 41. Compton, S.W., and Brownlee, R.G. BioTechniques, 6, 432 (1988). 42. Bocek, P., and Chrambach, A. Electrophoresis, 13, 31 (1992). 43. Hjerten, S. J. Chromatogr., 347, 191 (1985). 44. Ronai, Z., Guttman, A., Nemoda, Z., Staub, M., Kalasz, H., SasvariSzekely, M. Electrophoresis 2000, 21, 2058. 45. Stanchfield, J.E. and Batey, D.W. Poster Presentation at Genome Mapping and Sequencing Symposium, Cold Spring Harbor, NY, May, 13– 17, (1998), p. 214.

8 Development of Battery-Powered, Portable Instrumentation for Rapid PCR Analysis Phillip Belgrader and M. Allen Northrup Microfluidic Systems Inc., Berkeley, California

Bill Benett, Dean Hadley, Ray Mariella, Jr., Jim Richards, Paul Stratton, Shanavaz Nasarabadi, and Fred Milanovich Lawrence Livermore National Laboratory, Livermore, California

8.1 INTRODUCTION The explosive growth of biotechnology during the past decade will change the fundamental nature of many aspects of modern civilization. New drugs designed and manufactured using recombinant DNA technologies have benefited millions of people, from diabetics to those receiving genetically engineered vaccines. Gene therapy techniques may provide even more spectacular health benefits by permitting the body to cure itself; and engineered biological agents are being used for pest control and environmental restoration, just to name a few examples. This growth prompted, and was in turn fueled in part by, advances in bioengineering capabilities. These advances met the researchers call for faster and faster analyses and highly parallel measurement capabilities. One such analysis technique is the polymerase chain reaction 183

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(PCR) [1,2], a powerful method for the amplification, detection, and characterization of distinct nucleic acid regions within the genome of animals, plants, fungi, and microbes. PCR is widely used for applications such as human identification, infectious disease testing, and genetic analysis because of the small sample requirement, high specificity and sensitivity, short reaction time, simple setup, and robust nature of the reaction. The reaction is accomplished by adding a master mix (Taq DNA polymerase, MgCl 2, a pair of oligonucleotide primers, deoxynucleotides, buffer) and sample material containing target DNA, serving as a template, to a small reaction tube (25–100 µL). The tube is placed in a thermal cycling instrument that cycles the temperature of the reaction between two (or three) setpoint temperatures. A typical thermal cycle consists of a denaturing step at 95°C and an anneal/ extend step at 60°C (Fig. 1). At each cycle, the strands of the doublestranded DNA molecules are separated during the denaturing step, then replicated by the thermostable Taq DNA polymerase during the anneal/ extend step. After 30 cycles, at least 10 9 copies of a specific sequence region (50–1000 bp in length), termed an amplicon, are generated from each starting molecule of target DNA in the sample material. The efficiency of the PCR amplification is highly dependent on

Figure 1 Overview of PCR amplification. A single tube containing reagents and sample (i.e., bacteria) is placed in a thermal cycler. The thermal cycler continuously changes the temperature between 95°C and 60°C. At 95°C, the double-stranded DNA molecules are denatured into separate strands. At 60°C, a short synthetic oligonucleotide primer anneals to each strand, and the primers are extended by Taq DNA polymerase to replicate the sequence region between the primers. Since both strands of the DNA molecules are duplicated after each cycle, amplification is exponential, and after 1–2 hr in a conventional thermal cycler, billions of copies (amplicons) of a specific region of the genome are generated.

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the performance of the thermal cycler. Factors such as the heating and cooling rates and temperature accuracy and uniformity can effect the yield of PCR product. Currently, there are many brands and types of commercial thermal cyclers available, but when the microchip PCR project was initiated, the Perkin-Elmer GeneAmp PCR System 9600 (introduced in 1990) was the workhorse, and is still highly utilized today. The 9600 has an aluminum block with wells for 96 polypropylene reaction tubes. Heating and cooling is accomplished with a heater below the block and coolant from a refrigeration unit flowing through the block. A typical PCR run of 25–40 cycles takes 1–2 hr to generate a sufficient number of amplicons for subsequent analysis by agarose or polyacrylamide slab gel electrophoresis, capillary electrophoresis, microtiter plate ELISA assays, or oligonucleotide hybridization strips or arrays. Total time of a PCR amplification and detection can span ⬃2–6 hr. Between 1991 and 1993 Roche and Perkin-Elmer reported on a new PCR assay called Taqman [3–5] (Fig. 2). Taqman is similar to traditional PCR except that the reaction contains a fluorescence resonant energy transfer probe (FRET). The probe typically contains a green fluorescence dye (6-Fam), termed the reporter, at one end and an orange fluorescence dye (Tamra), termed the quencher, at the opposite end. The close proximity of the dyes causes the quencher to suppress (or absorb) fluorescence emission by the reporter. As probe hybridizes to the amplicons during PCR, it is cleaved by the nuclease activity of the Taq DNA polymerase. The reporter is liberated from the effects of the quencher, and the resultant increase in green fluorescence emission can be monitored with an external fluorescence detector. The amount of fluorescence is proportional to the number of amplicons generated in the reaction. In 1995, Perkin-Elmer introduced the ABI Prism 7700 spectrofluorometric thermal cycler for real-time Taqman analysis [6]. This instrument utilizes a 9600 thermal cycler with a modified lid that contains fiber optics terminating above each reaction tube. The fibers simultaneously direct an argon ion laser into each tube and collect the full-spectral fluorescence emissions. The emitted light passes through a series of lenses and filters, and images the light on a charged-couple device (CCD) camera. The advantages of this system include (1) simul-

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Figure 2 The Taqman reaction. A fluorescence resonance energy transfer (FRET) probe is present in the PCR reaction. As PCR product is produced, the probe hybridizes to a complementary strand during the anneal/extend step at 60°C. The probe is cleaved and displaced by the Taq DNA polymerase as it extends a primer hybridized downstream on the same strand. This cleavage allows the reporter fluorescence dye to become liberated from the effects of the quencher fluorescence dye. The increase in the reporter fluorescence signal emitted is measured during each cycle.

taneous PCR amplification and detection, (2) closed tube format to prevent cross-contamination, and (3) real-time data acquisition for quantitative measurements based on reaction kinetics. An area of recent, rapidly growing national-security concern is the potential for biological terrorism. Given that relatively small quantities of agent can potentially contaminate hundreds of square kilometers, several nations without a nuclear deterrent but with a modern biological capability have chosen to pursue biological warfare (BW) as a less effective, but more affordable method of protecting themselves against better-armed adversaries. Biological weapon capability can also be

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readily acquired by subnational and terrorist groups and its potential use in terrorist activities is considered a major rising threat to all civilian populations. As part of an overall strategy to respond to this threat there is an urgent need to equip emergency first responders with the capability to rapidly and accurately identify biological agents in the field. Despite the growth in PCR applications, a truly portable PCR capability for environmental and point-of-use testing for pathogenic microorganisms had been nonexistent because of size, weight, analysis time, and power requirements of available instruments. For example, the Prism 7700 weighs 342 lb, operates on 2600 watts, and takes ⬃2 hr to complete a run. This chapter will summarize years of effort at Lawrence Livermore National Laboratory, motivated at first by the need for faster diagnostics in support of biotechnology developments and recently by the need for counterterrorism tools, to design, build, and test rapid, portable PCR instruments for high-performance and real-time analyses in any location. 8.2 INSTRUMENTATION The microchip PCR instruments consist of a thermo-optic module(s), a set of electronic control PC boards, a laptop computer, customized software, a power supply, and a case. Two instruments that resulted from this project are shown in Figure 3. The MATCI [9] (miniature analytical thermal cycling instrument) was the first portable, batterypowered spectrofluorometric thermal cycler for real-time PCR. The MATCI fits in a briefcase, contains a single thermo-optics module, and is powered by 13 NiCad batteries. The reaction is monitored in real time, enabling a positive signal to be identified before the run is completed. Several reports have been published describing the MATCI and its applications to infectious disease detection and identification, genetic analysis, and human identity testing [9–11]. The next instrument, the ANAA [13,14] (advanced nucleic acid analyzer), contains an array of 10 thermo-optics modules, improved software, and a modified detection system. These features expanded capability, increased sensitivity, and simplified the user interface. The thermo-optic module, control boards, and software will be described in more detail.

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Figure 3 The MATCI (top) and ANAA (bottom). High-performance instruments for rapid real-time PCR (TaqMan) analysis. These instruments offer battery power operation, real-time monitoring of the reaction, and automatic positive calling. Since there are no moving parts (except the cooling fans), the instruments are rugged and quiet.

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8.2 Thermo-optic Module The thermo-optic module is the fundamental component of the instruments. It contains a micromachined silicon reaction chamber with an integrated heater, a temperature sensor, an air ducting system for cooling, and focusing and collecting optics for real-time detection. Reaction Chamber The first PCR microchip or chamber [7] was inspired by the growing field of MEMS (microelectromechanical systems) [12] in which the microfabrication techniques used to generate integrated circuits from silicon wafers were being applied to creating microdevices such as accelerometers and pressure sensors. Silicon was expected to be an ideal material for a thermal cycling chamber since (1) silicon was an excellent heat sink, with thermal conductivity only slightly less than that of aluminum; (2) undoped silicon is normally a poor enough electrical conductor, and can be directly coated with a metal or doped-polycrystalline-silicon film to function as an integrated resistive heater; (3) the chamber could be shaped to exhibit a high surface area to maximize the rate of heating and cooling and provide even heat distribution; and (4) the mass of the chamber could be relatively small, enabling the integrated heater to operate on low voltage. A crude PCR microchip, designated chamber I (Fig. 4a), was fabricated using standard masking, photolithography, and bulk anisotropic silicon etching in KOH with a silicon-nitride film for protection. There was no online detection capability since real-time fluorogenic assays had yet to be developed. The performance of this chamber was less than that of the standard commercial systems in terms of productivity per thermal cycle, with the ideal being a doubling of the concentration of the target DNA sequence on each cycle. The heating and cooling rates were very rapid, 10°C/sec or faster, but the uniformity of temperature throughout the reaction mixture was unacceptably large. Temperature measurements were made using small-diameter thermocouples, inserted into the solution, and via imaging on photometric IR cameras. One major problem was materials in contact with the PCR solution showed inhibition of the reaction. Even when coated with siloxanes that were compatible with PCR, cleaning a chamber after its use was inconvenient and manu-

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Figure 4 Evolution of the chamber for efficient heating and cooling of the reaction. (A) Chamber I, (B) chamber II, and (C) chamber III. The optical window was introduced in chamber II for real-time fluorescence detection.

ally intensive; several improvements were necessary. These considerations led, ultimately, to the sleeve designs with a plastic insert to contain the PCR solution. The early flat designs had originally been selected since they offered, from an engineering perspective, the most power-efficient manner in which to heat and cool the PCR solution—heating a flat, thin sample could be effected easily using a very-low-mass thin-film resistive heater on a silicon nitride window, and cooling such a shape

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was also straightforward. Unfortunately, such a design led to variations up to 10°C in solution temperature across its flat dimensions. Therefore, a hexagonal chamber, designated chamber II (Fig. 4B), was prepared by bonding two identical halves of etched silicon. The hexagonal chamber results from the anisotropic etching of the (100) face of the singlecrystal silicon. A thin film of metal or doped polysilicon served as the heater. The relative dispersion of heat, developed by the thin-film heater, between the silicon and the surrounding air is dependent upon the effective thermal transport properties of each. Since during heating there is no forced convection of the air, the much larger thermal conductivity of the silicon means that nearly all of the heat is deposited into the silicon. An optical window was positioned at the center of the chamber to accommodate an integrated fluorescence detector, described below. Initially, a thin layer of silicon nitride covered the window to retain the PCR mix that was added to the chamber. However, tests had determined that the uncoated silicon surface inhibited PCR amplification, and therefore, a special disposable polypropylene tube and chamber with an open window were produced [8]. There was some reduction in the transfer of heat through the plastic (heating and cooling rates were decreased by about 2°C/sec), but the use of the plastic insert increased the processing yield and decreased the time that was required to fabricate the chambers, since they did not need the silicon nitride window nor internal passivating coatings. Also, the use of the plastic insert greatly increased the number of samples that could be processed in a chamber per day, since it was no longer necessary to clean the inside of the chamber after each use. Both the MATCI and the ANAA use chamber II. Temperature Sensor Electrical contact is made to the chamber by the deposition of two gold pads on each of the two heaters (Fig. 4B). The interior walls of the chamber module contain circuit boards that carry current to the heaters from a connector on the rear of the board (Fig. 5). Connection from the board to the chamber is made by beryllium copper spring fingers that touch each heater at only four points, thereby adding minimal thermal mass to the system. The temperature of the chamber is monitored

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Figure 5 Diagram of the thermistor placement and the logical feedback temperature set and control circuitry.

by means of either a thermocouple (used in the MATCI) attached to the chamber wall or a thermistor probe (used in the ANAA) inserted into the lower end of the chamber. Contact is made to the thermistor by a second set of thin metal fingers that carry the signal to a custom circuit board. Air Ducting System Cooling is accomplished by forcing ambient air past the chamber during the cooling steps. For the ANAA, this is accomplished by two fans pulling cool air through manifolds located below each bank of five thermo-optic modules. The MATCI has one fan. The exterior walls of the module have machined passages that serve as air ducts. Air is pulled through openings at the top of the module, passed through the exterior wall ducts, and injected directly onto the heated faces of the chamber. The air then moves through ducts formed by the interior walls of the module, out openings in the bottom of the module, and into the manifold. Focusing and Collecting Optics The detection module in the MATCI is shown in Figure 6. The inexpensive detection components consist of an excitation source, focusing and turning optics, filters, and detectors. The excitation source, located in

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Figure 6 Illustration of the MATCI detection system.

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the module cover, is a blue LED which peaks near 490 nm, followed by a bandpass filter to minimize any direct contribution from the LED to the detectors. The excitation light is transmitted down the reaction tube. Fluorescence from the PCR sample is collected through two collimating ports that flank the two sides of the 1-mm ⫻ 2-mm window in the wall of the silicon chamber. The two signals are filtered with bandpass filters that transmit fluorescence wavelengths but reject excitation wavelengths. A piece of absorbing colored glass eliminates any residual excitation light that may exist before the signals reach their respective detectors. The detectors are standard silicon photodiodes serving as charge integrators, and require 1–5 sec to collect the fluorescence emissions from the chamber. The optic system was modified for the ANAA as shown in Figure 7. The excitation source is illuminated from the side instead of the top. The excitation light passes through a focusing port to reach the sample via the window in the chamber wall. Fluorescence from the PCR sample is collected through a collimating port directly opposite the excitation port, then illuminated on mirrored right-angle prisms which reflect

Figure 7 Diagram of the ANAA detection system.

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the beam to the detectors. This iteration provides 10 times greater sensitivity, faster operation (0.1 sec collection time), better signal-to-noise ratio, and greater sensitivity to weak target concentrations. In addition, the side illumination allows free access to the top of the chambers, which is essential for integration of automated sample preparation. Other improvements to the detection system include an optimized LED, removable optical filters, better electrical shielding, and a more compact, modular design. 8.2.2

Control Boards

There are four types of custom PC boards: the distribution board, the motherboard, the heater control board, and the detector board. The distribution board buffers and distributes signals and power between the laptop computer, the power system, and the motherboard(s). It also controls the PCR cooling fans. The motherboard(s) distributes the signals and power between the heater control boards, detector boards, and the distribution board. The ANAA contains 10 heater control boards that (1) control the heat delivered to the individual thermal chambers and the 12 detector boards, (2) measure the fluorescence in the chambers, and (3) convert the optical signals into electronic signals. 8.2.3

Software

The software application IGOR Pro (WaveMetrics, Lake Oswego, OR) was chosen during the development of the MATCI. IGOR specializes in waveform analysis and is capable of interacting with external hardware through a National Instruments data acquisition card. The MATCI uses a single PCMCIA card to control the thermal system and the detection system. The ANAA has two of these cards. The four major sections of the user interface software for the PCR instruments are setup, run displays, calibration, and data management. For the MATCI, setup is limited to the setting of PCR temperatures, temperature hold times, and detection integration (fluorescence collection) times. Values are entered by directly editing the software code. Run displays on the computer monitor (Fig. 8) include a ‘‘strip chart’’ depiction of the cycling temperature and a real-time signal display. When a positive sample is detected, the user is notified via an audible beep and a screen message. A custom algorithm, based on filtered slope

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Figure 8 Run display on the laptop computer in the MATCI (top) and the ANAA (bottom).

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analysis that discriminates against noise, determines when a sample is positive. Thermal calibration on the MATCI utilizes an external thermocouple to monitor the sample temperature while the heater is set to various setpoints. The user records calibration values and utilizes builtin IGOR functions to curve fit the thermal response of the system. Run data is archived by saving the entire experiment. PCR run data management and analysis require the user to be familiar with IGOR. The basic functionality of the control system for the ANAA is similar to the MATCI, with some additional features. A continuous run mode function allows the user to stop and start a particular chamber without affecting other chambers that are running. The setup utilizes dedicated panels, which prompt the user for information while protecting the software code from accidental corruption. Checks and guides ensure that the user enters reasonable values. In addition to entering PCR parameters and detection variables, the ANAA also supported assigning the various chambers with identifying labels. This is important when several types of assays are used in a single run or when a chamber is used in 2-plex mode (two Taqman probes). When starting a new run, the user is prompted for a run ID, which is checked for uniqueness to avoid overwriting existing files. Run displays on the ANAA (Fig. 8) include a dedicated graph for each of the 10 chambers, a graph showing initial signal levels, a chamber assignments window, the IGOR control/history window, and an ANAA main panel. The main panel indicates run ID, cycler status, detection status, and the time since the run started. The chamber assignments window has a text box that indicates the particular assay run in each chamber. The box changes color from green to red when a positive is detected. The detection algorithm is essentially the same as that for the MATCI, but the filter is more stringent due to the reduced background signal. Calibration on the ANAA is partially automated and fully documented to ensure accuracy and efficiency. The user is prompted to move the thermocouple from chamber to chamber and enter external liquid thermocouple readings. A complete on-line manual discussing calibration as well as other topics for the ANAA is incorporated into the IGOR Help reference system. In addition to calibration, several other maintenance routines are available that to tune up and completely characterize ANAA performance.

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Data management on the ANAA includes automatic data storage of two files. One file contains all pertinent setup and result information while the other contains only the key results from the run. Both are supported by separate IGOR programs designed to fully analyze and display the data. IGOR allows the user to simultaneously compare results from different runs. The information from each run is clearly labeled, complete with assay types and run ID displayed on graph legends. 8.3

PERFORMANCE

The ANAA contains an array of 10 modules, and each module can be independently stopped and started without effecting the other modules. Thus, each thermo-optic module acts as an independent real-time PCR instrument. An important metric for system performance was the ability to acquire similar real-time detection signals among the array of modules. Figure 9 shows the analysis at 38 sec/cycle of replicate samples of Bacillus anthracis DNA in modules 1–9. A TaqMan probe was used that identifies plasmid pX02, a pathogenic marker. Module 10 served as the negative control. Two important real-time detection signal characteristics are assessed: the threshold cycle, the cycle when a significant increase in a positive fluorescence signal amplitude is measured above background, and the endpoint value, the signal amplitude at the last cycle. The threshold cycle (CT) inversely correlates with the initial quantity, or copies, of target DNA molecules in the reaction. In this run, the threshold cycle for modules 1–9 was 22. Typically, a log increase in the initial quantity of target, decreases the CT by 3–4 cycles. The endpoint value also correlates with the quantity of target, but should be considered to be semiquantitative. The endpoint value typically exhibits more variation since the reaction often becomes saturated with PCR product and depleted of reactants (e.g., primers, nucleotides) and active enzyme by this stage. Nevertheless, modules 1–9 displayed similar endpoint values. Another run demonstrated the quantitative capability of the ANAA. Ten-fold serial dilutions of the B. anthracis DNA were simultaneously analyzed at 27 sec/cycle using a TaqMan probe specific for plasmid pX01, another pathogenic marker. The data in Figure 9 indi-

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Figure 9 Fast detection of Bacillus anthracis. (Upper panel) Reactions containing 10 pg of B. anthracis genomic DNA were analyzed in modules 1–9 of the ANAA at a rate of 38 sec/cycle. Module 10 (no template) contained a reaction without genomic DNA. The probe recognized the CAP gene in plasmid pX02. (Bottom panel) Serial 10-fold dilutions of the B. anthracis genomic DNA were analyzed at a rate of 27 sec/cycle using a probe that recognized the PA gene in plasmid pX01. The amount of DNA in each reaction is indicated.

cated that the amount of starting DNA in the reaction was inversely correlated to the threshold cycle. A detection limit of 100 attograms, or 20 copies, was observed. Further optimization of the ANAA enabled the cycle time to be reduced to as little as 17 sec [14]. At this ultrafast speed, the efficiency of the reaction was still maintained, and the pro-

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Figure 10 Ultrafast detection of Yersinia pestis cells. Reactions containing 500, 50, and 5 cells were analyzed at a rate of 17 sec/cycle.

ductivity of amplicons during each cycle was not compromised. Figure 10 shows ultrafast detection of Yersinia pestis cells. Cell lysis, PCR, and detection of 500 Y. pestis cells was accomplished in only 8 min. The assay was still quantitative, with a limit of detection of 50 cells. These results clearly demonstrate that the modules exhibit similar thermal and detection characteristics. More importantly, the results should change our way of thinking about DNA analysis. From what used to take days using Southern blot hybridization, to several hours using traditional PCR methods, and now to minutes using technology that is not only fast and efficient, but portable. 8.4

NEXT-GENERATION INSTRUMENT

This work has resulted in the progressive size reduction of the thermooptics module while decreasing the time required for real-time PCR

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Figure 11 The HANAA. The first hand-held, battery-powered, real-time PCR detection instrument. The battery pack, shown on the left, clips onto the user’s belt and provides power for 2 hr of continuous use (⬃10 runs).

analysis. The next-generation instrument, called HANAA (hand-held nucleic acid analyzer) has recently been built (Figure 11). HANAA contains four thermo-optic modules, an integrated Pentium processor, a keypad, and an LCD. The modules, half the size of those used in the ANAA, utilize a new chamber (chamber III) and a more compact optical configuration that eliminates the prisms. Two optical windows accommodate both blue and green LEDs. Chamber III (Fig. 4C) has two vertically oriented, anisotropically etched V-grooves to the planar face of the halves. These grooves remove thermal mass and also help to thermally isolate the central region that holds the tube. The grooves also create additional surfaces for resistive heating and air-cooling. The applied heater current is heating much less material and heating the sample much more directly than in the previous design. A second modification is the addition of a series etched V-grooves on the opposite planar face of the silicon halves. These grooves run perpendicular to, and intersect with, the vertical Vgrooves. When the silicon halves are connected, a series of short channels are created that provide a path for air to be forced directly past

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Figure 12 Comparison of the thermal performance of chamber II and chamber III. Setpoint temperatures are indicated. Chamber III can heat and cool the solution faster than chamber II. The overshoot programmed for the chamber temperature allows the solution to be heated at a fast rate.

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Figure 13 Ultrafast detection of Erwina herbicola cells using the HANAA module. (Upper panel) Thermal profile of the reaction. (Lower panel) Realtime detection profile. Five hundred cells were detected in only 6 min using a thermal cycle rate of 14 sec/cycle.

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the surfaces of the thermally cycled, central region of the device. A third modification is the replacement of the doped polysilicon with platinum as the thin-film resistive heater. Platinum has a stable coefficient of thermal resistance that allows it to be used as both a heater and a temperature sensor. It is a well-known principle of control engineering that one can attain the tightest temperature control of an object if the temperature sensor is placed directly on the heater itself. The combination of lower overall thermal mass and better temperature control led to the improved performance results shown in Figure 12. The control software is located with dedicated onboard microcontrollers and therefore separated from the user interface software. The keypad and LCD guide the user through a series of menus to set up and run the instrument. The LCD displays bar graphs representing the fluorescence signal intensities. The total power requirement for HANAA, including the computer, is only 40 watts. Power for 2 hr of continuous operation is provided by a 1-lb external battery. Calibration is completely automated for each chamber. The performance of a HANAA module is shown in Figure 13. Five hundred bacteria cells were subjected to real-time PCR analysis. The thermal and detection profiles demonstrate 14 sec/cycle and 6-min detection. 8.5

CONCLUSION

The technology described represents years of effort dedicated to building portable DNA analytical instruments that perform better than traditional bench top systems. This was accomplished by substituting and integrating miniature, inexpensive, low-power components in place of bulky, high-power components. The potential applications for portable DNA analytical instruments are enormous and should provide exciting new capabilities to allow first responders, soldiers, clinicians, forensic scientists, environmentalists, epidemeologists, and food inspectors to perform rapid on-the-spot testing for pathogenic microbes in any location. In addition, this new paradigm of performing nucleic acid–based microbial detection within minutes using portable, battery-powered instruments is certain to generate applications not conceived of before because development of this technology was not considered possible.

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ACKNOWLEDGMENTS This document was prepared as an account of work sponsored by an agency or agencies of the U.S. Government. Neither the U.S. Government nor the University of California nor any of their employees make any warranty, express or implied, or assume any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represent that its use would not infringe privately owned rights. Reference herein to any specific commercial products, process, or service by trade name, trademark, manufacturer, or otherwise does not necessarily constitute or imply its endorsement, recommendation, or favoring by the U.S. Government or the University of California. The views and opinions of authors expressed herein do not necessarily state or reflect those of the U.S. Government or the University of California, and shall not be used for advertising or product endorsement purposes. This worked was supported and funded by the Department of Energy, Defense Advanced Research Projects Agency, and Central MASINT Organization and was performed under the auspices of the U.S. DOE by Lawrence Livermore National Laboratory, Livermore, CA, under contract No. W-7405-Eng-48. We also thank Vicky Hambrick for assisting us with this work.

REFERENCES 1. Saiki, R., Scharf, S., Faloona, F., Mullis, K, Horn, G., Erlich, H., Arnheim, N. (1985) Enzymatic amplification of beta-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230:1350–1354. 2. Mullis, K., Faloona, F., Scharf, F., Saiki, R., Erlich, H. (1986) Specific enzymatic amplification of DNA in vitro: the polymerase chain reaction. Cold Spring Harbor Symp. Quant. Biol. 51:263–273. 3. Holland, P.M., Abramson, R.D., Watson, R., Will, S., Gelfand, D.H. (1991) Detection of specific polymerase chain-reaction product by utilizing the 5′-3′ exonuclease activity of Thermus aquaticus DNA-polymerase. Proc. Natl. Acad. Sci. USA 88:7276–7280. 4. Livak, K.J., Flood, S.J.A., Marmaro, J., Guisti, W., Deetz, K. (1995)

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9 Practical Aspects of Sheath Flow Fraction Collection in Capillary Electrophoresis Odilo Mu¨ller Agilent Technologies Deutschland GmbH, Waldbronn, Germany

Hirofumi Suzuki Organon, Ltd., Osaka, Japan

9.1 INTRODUCTION Capillary electrophoresis (CE) has recently become a widely used method for the analysis of a large variety of substances including small ions, chiral substances, and biomolecules. In many cases (e.g., analysis of small molecules), separation of the sample constituents may be adequate. In many other cases (e.g., separation of biomolecules), however, isolation (⫽ fraction collection) and further characterization of the separated bands would be desirable if not mandatory. Classical separation techniques used in the field of bioanalysis (e.g., HPLC and slab gel electrophoresis) offer not only the option of separating but also of collecting the separated bands. CE, on the other hand, which offers superior separation performance (efficiency, speed of separation) for many applications, presents several challenges for its preparative operation: only minute sample amounts can be loaded onto the CE column and the CE separation requires the presence of a closed electrical circuit, making it difficult to collect the eluting bands. Furthermore, each fraction collection technique for CE presents its own problems. 207

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There are several ways to collect fractions from CE systems. The most widely used technique is electroelution [1], where the electrical field is switched off when a detected band reaches the end of the capillary. After transferring the capillary outlet from the buffer vial to a collection vial containing a small amount of buffer (⬃10 or more µL), the field is switched on again to allow the band to migrate into the collection vial. After this collection step the electrical field is switched off again, the capillary outlet is transferred back to the buffer vial, and the run is completed with the electric field applied. Pressure elution [2] can be performed analogically with the sole exception that for collection a pressure (instead of the electrical field) is applied. Electroelution and pressure elution can be performed on commercially available instrumentation. They suffer from the fact that they are discontinuous collection techniques, which means that the electrical field is switched off for collection. The predicted elution time of the detected bands becomes, therefore, increasingly inaccurate with the number of collected bands. It is highly impractical to collect more than three to five bands in a single run using these approaches. Continuous collection techniques (electrical field remains on during collection) allow equally accurate collection of all separated bands. Those techniques are wetted membrane collection [3], collection from frit structures [4], and sheath flow collection [5]. In the case of wetted membrane collection the elution side of the capillary stays in permanent contact with a membrane which is soaked with the separation buffer. If the membrane is moving (e.g., rotating), the eluting species are immobilized on the surface in the order of their mobilities. Frit structures, on the other hand, allow the closing of the electric circuit before the end of the capillary by means of a porous connection which is surrounded by buffer. A strong electro-osmotic flow (EOF) can allow to transport the bands beyond this connection point so that they can be collected at the end of the capillary column. Both wetted membrane collection and collection from frit structures are limited since, in the first case, the bands stick to a solid surface and have to be eluted for many further applications; in the second case, an EOF is required for elution from the capillary, which is not desirable for many separations of biomolecules (polymer matrices).

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Sheath flow collection is the most versatile continuous collection technique. In principle, it allows accurate collection of an unlimited number of bands. The collected species are directly available in solution for further analysis. In this approach, the capillary is inserted into a T-connection (sheath flow connection), which is sealed to one side by means of a septum (Fig. 1). On one side of the T-connection a Teflon tube is glued to serve as collection tip. Buffer, which is provided by a syringe pump, is forced to flow out of the Teflon tip, as the other side of the T-connection is sealed by means of a septum. A closed electric circuit is obtained if the capillary is connected to the sheath flow cell on one side and inserted into a buffer vial on the other side, and a power supply is connected to both the sheath buffer and the buffer reservoir. Species that migrate lower than the flow velocity of the sheath liquid inside the Teflon tip stay in the droplet that builds at

Figure 1 Schematic view of sheath flow cell including holder for optical fibers. Insert: Teflon collection tip with collection buffer droplet.

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the end of the Teflon tip (Fig. 1, insert) and can be collected without the need of switching the voltage off (see Note 1). For the practical work with a sheath flow fraction collector (as well as with any other fraction collector), two major points have to be taken into consideration: the small sample amounts (see Note 2) and the collection precision. To increase the collected sample amount two measures can be taken: employment of larger capillary inner diameters (ID) or pooling of collected samples from several collection runs. Of course the use of a larger capillary ID is limited since at a larger ID band broadening will occur. With regard to the capillary diameter the rule of thumb is to employ the biggest ID possible which results in the required resolution. Pooling of collected samples compromises the advantage of the speed of CE separations and increases the risk of sample contamination (one incorrectly collected sample contaminates the rest of the pool). There is, however, an increasing number of analytical techniques which is able to analyze the small sample amounts collected from CE separations. Among those techniques, mass spectrometry, which is evolving very rapidly in recent years, plays a major role due to its ability to reveal mass information as well as structural information. Other useful techniques for further analysis of the collected species are microsequencing, polymerase chain reaction (PCR), and immunoassays. Two CE separation modes are especially suited for fraction collection: capillary isoelectric focusing (cIEF) and DNA applications. In the case of cIEF separations, the entire capillary is filled with the ampholyte/sample mixture, thus increasing the overall sample amount. DNA applications have the unique advantage that PCR can be used for sample amplification prior to secondary analysis such as cloning, sequencing, or mutation detection. Collection precision is another important parameter in CE fraction collection. A major advantage of CE consists in the fact that it is a high-resolution technique. Therefore, it is important to maintain the resolution during the collection process. Several measures can help to improve collection precision. The first step consists in choosing a continuous collection technique (e.g., sheath flow collection) as discussed above. In addition it is important to place the detection point as close to the capillary exit as possible. This measure ensures exact prediction

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Figure 2 Two collection modes: peak activated collection (PAC) and cutting band collection (CBC).

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of the elution time of a detected band (see Note 3). One way of placing the detection point close to the capillary exit is the extension of the light path by means of optical fibers. Optical fibers allow detection as close as a few millimeters from the capillary outlet, thus ensuring a high collection precision. Collection precision is also improved by automating the collection procedure. A computer monitoring the detection signal can be programmed to collect all detected bands into appropriate collection vessels. Such collection vessels can be glass capillaries which minimize sample evaporation and facilitate the handling of small sample amounts. There are two general ways of collection: peak activated collection (PAC) and cutting band collection (CBC) (Fig. 2). For PAC a threshold is set to define a peak. All time values exceeding the threshold indicate a band passing the detector. The beginning and end of this band (t 1 and t 2; collection window) are used for calculation of the elution time of this band. According to the application it might be useful to vary the size of the collection window (t 1 ⫺ x, t 2 ⫹ x) or account for a systematic shift (offset). PAC is used for collection of baseline resolved bands. If unresolved bands are encountered, CBC is advantageous. It allows definition of a collection window to cut the unresolved

Figure 3 Sheath flow fraction collector.

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bands into small portions. The beginning of the unresolved band is again recognized by a threshold and the use of an offset value ensures complete collection of the band. In this approach, each collected fraction has to be examined individually to verify purity of the collected species. Figure 3 shows a fraction collector in which a sheath flow unit, optical fibers, and computer-controlled autosampler for glass capillaries have been implemented. This fraction collector allows accurate collection of up to 60 fractions (number of collection capillaries that can be installed on the holder). In the following, collection procedures for three general CE modes (separation in polymer matrices, cIEF, and CZE) are presented. 9.2 MATERIALS 9.2.1

Calculation of Back-Flow Velocity

1. Sheath flow fraction collector equipped with optical fiber detector, collection capillaries, stepper motor, syringe pump, and appropriate computer control (see Fig. 3). 2. Separation capillary, which is used for subsequent experiments. 3. Separation buffer (40 mM Tris/TAPS, ampholyte/H 2O mixture (2%) or 10 mM ammonium acetate) (see Note 4). 4. Fluorescein: 10 ⫺4 M in destilled H 2 O. 9.2.2

DNA Fraction Collection from Polymer Filled Columns

Collection 1. Instrument from Sec. 9.2.1. 2. Polysiloxane coated capillary (J&W, DB-1, 100 µm ID, L ⫽ 30 cm, 1 ⫽ 29 cm). 3. DNA sample (10–50 µg/mL). 4. Size exclusion spin columns for desalting of DNA (see Note 5). 5. 40 mM Tris/TAPS buffer (pH 8.4) in deionized water. 6. Methyl cellulose (MC): 1% (w/v) in 40 mM Tris/TAPS buffer.

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Purity Check of Collected DNA Samples (See Note 6) 1. Instrument equipped with laser induced fluorescence detector (excitation wavelength: 488 nm, emission of DNA/EtBr-complex: 605 nm). 2. Polysiloxane coated capillary (J&W, DB-1, 50 µm ID, L ⫽ 20 cm, 1 ⫽ 12 cm). 3. Chemicals from Sec. 9.2.2. 4. Ethidium bromide (EtBr): 1 µg/mL in 1% MC matrix (see Note 7). 9.2.3

Protein Fraction Collection from cIEF Separations

Collection 1. Instrument from Sec. 9.2.1. 2. Polyvinyl alcohol (PVA) coated capillary (75 µm ID, L ⫽ 40 cm, l ⫽ 39 cm). 3. Fluorescein: 10 ⫺4 M in deionized water. 4. Ampholyte: 1% (1:1:1, v/v) mixture of Pharmalyte, Ampholine, Servalyte in deionized water. 5. Proteins: 0.05–0.1 mg/mL in ampholyte mixture (see Note 8). 6. Acetic acid: 0.5% (v/v) in deionized water. 7. Ammonium hydroxide: 0.3% (v/v) in deionized water. MALDI-TOF Mass Determination 1. MALDI/TOF mass spectrometer. 2. Trifluoroacetic acid (TFA). 3. Acetonitrile. 4. Sinapinic acid: 0.1 M in TFA/acetonitrile (60:40 v/v) 9.2.4

Sugar Fraction Collection from CZE Separations

Collection 1. Instrument from Sec. 9.2.1. 2. Polyvinyl alcohol (PVA) coated capillary (75 µm ID, L ⫽ 30 cm, l ⫽ 29 cm).

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3. 1-Aminopyrene-3,6,8-trisulfonate (APTS): 0.2 M in 15% acetic acid. 4. Sodium cyanoborohydride: 1 M in tetrahydrofuran (THF). 5. Size exclusion spin columns. 6. Ammonium acetate: 10 mM in deionized water, pH 4.75. MALDI-TOF Mass Determination 1. MALDI/TOF mass spectrometer. 2. 6-Hydroxypicolinic acid (6-HPA): 4 mg/mL in acetonitrile/ H 2O (7:3, v/v). 3. 3-Hydroxypicolinic acid (3-HPA): 40 mg/mL in acetonitrile/ H 2O (7:3, v/v). 4. Cation exchange resin (NH 4 ⫹ form) [6] 9.3 METHODS 9.3.1

Calculation of Back-Flow Velocity (see Note 9)

1. Place separation capillary in sheath flow connection (see Note 10). 2. Select flow rate of sheath flow buffer (usually between 8 and 12 µL/min). 3. Fill separation capillary with fluorescein solution (10 ⫺4M). 4. Insert separation capillary into buffer vial and mark this moment on chart recorder (see Note 11). 5. Measure time between beginning of the backflow (capillary insertion) and replacement of the fluorescein solution by the sheath buffer in the detection window (see Note 12). 6. Calculate backflow velocity. 7. Adjust the height of the inlet buffer vial for all collections to counterbalance the pressure created by the sheath liquid (see Note 13). 9.3.2

DNA Fraction Collection from Polymer Filled Columns

Collection 1. Dissolve 1% methyl cellulose (MC, 2% ⬅ 4000 cps) in 40 mM Tris/TAPS buffer by gently stirring the mixture overnight.

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2. Degas solution in mild vacuum, shake if bubbles are formed, and slowly increase vacuum. This may take 1/2 hr (see Note 14). 3. Desalt sample according to operating instructions of spin columns. 4. Insert separation capillary in sheath flow connection. 5. Fill separation capillary with methyl cellulose solution and wipe Teflon tip clean (see Note 15). 6. Position a collection capillaries close to Teflon tip (distance ⬇ 300 µm). 7. Separate DNA samples according to the following protocol: Capillary: 100 µm ID capillary polysiloxane coated. L ⫽ 30 cm, l ⫽ 29 cm. Separation matrix: 1% MC, 40 mM Tris/TAPS. Injection: electrokinetically, 3 sec @ 400 V/cm. Electric field strength: 400–600 V/cm. Flow rate of sheath buffer: 12 µL/min. Detection: UV 254 nm. 8. Collect according to PAC or CBC protocol, depending on the resolution of the bands (see Note 16). 9. Blow collected species out of collection capillary into appropriate vessel for purity check, PCR, sequencing, cloning, mutation detection, or other applications.

Purity Check of Collected DNA Samples (See Note 17) 1. Perform CE separation in entangled polymer solution using the following conditions: Capillary: 50 µm ID capillary polysiloxane coated, L ⫽ 20 cm, l ⫽ 12 cm. Separation matrix: 1% MC, 40 mM Tris/TAPS. Injection: electrokinetically, 5 sec @ 400 V/cm. Electric field strength: 400–600 V/cm. Two buffer vials. LIF detection of EtBr stained DNA fragments: (488 nm/600 nm) (see Note 18). 2. Assess purity of sample.

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Protein Fraction Collection from cIEF Separations

Collection 1. Insert separation capillary and adjust height according to Sec. 9.3.1 (see Note 19). 2. Measure mobilization flow velocity by dipping capillary inlet into fluorescein solution and lift inlet like during mobilization (⬃4 cm; see Note 20). 3. Enter value of mobilization flow velocity into software (see Note 21). 4. Fill separation capillary with ampholyte/sample mixture. 5. Perform focusing according to the following protocol: Capillary: 75 µm ID capillary PVA coated. L ⫽ 40 cm, l ⫽ 39 cm. Separation matrix: 2% ampholyte mixture in H 2O. Anolyte: 0.5% (v/v) acetic acid. Catholyte: 0.3% (v/v) ammonium hydroxide (⫽ sheath buffer!). Injection: flush capillary with sample/ampholyte mixture. Electric field strength: 10 kV for 2 min and 30 kV for 13 min. Flow rate of sheath buffer: 12 µL/min. Detection: UV 280 nm. 6. Mobilize focused zones by lifting the inlet buffer vial for ⬃4 cm (the electric field remains on). 7. Collect fractions using PAC or CBC (compare Sec. 9.3.2). 8. Analyze isolated protein fractions by microsequencing, tryptic digestion, or MS analysis. MALDI-TOF Mass Determination 1. Prepare fresh MALDI matrix (saturated solution of sinapinic acid in 0.1 M trifluoroacetic acid (TFA)/acetonitrile (60:40, v/v). 2. Apply 2 µL collected protein onto MALDI target. 3. Wait until solvent evaporates. 4. Add 2 µL MALDI matrix solution. 5. Operate MALDI/TOF instrument according to instrumental instructions.

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Alternatively: 6. Prepare fresh MALDI matrix. 7. Dry protein fractions in speed vac. 8. Add 2 µL MALDI matrix solution and apply to MALDI target. 9. Operate MALDI/TOF instrument according to instrumental instructions. 9.3.4

Sugar Fraction Collection from CZE Separations

Collection 1. Insert separation capillary and adjust height according to Sec. 9.3.1. 2. Derivatize sugars according following protocol: Add 2 µL of 0.2 M APTS solution in 15% (v/v) acetic acid with 2µL of 1 M NaBH 3CN in THF to 10–50 nmol dried sugar. Heat 16 hr at 37°C. Purify reaction mixture using size exclusion chromatography spin columns. 3. Place capillary in sheath flow cell and adjust collection capillaries. 4. Separate sugars according to the following protocol (see Note 22): Capillary: 75 µm ID capillary PVA coated. L ⫽ 30 cm, l ⫽ 29 cm. Separation matrix: 10 mM ammonium acetate (pH 4.75). Injection: 5 sec @ 400 V/cm. Electric field strength: 300–500 V/cm. Flow rate of sheath buffer: 12 µL/min. Detection: UV 254 nm. 5. Collect desired bands according to PAC or CBC method. 6. Analyze isolated sugars using MS or other techniques. MALDI-TOF Mass Determination 1. Prepare 6-hydroxypicolinic acid (6-HPA, 4 mg/mL) in mixture of acetonitrile/water (7:3, v/v).

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2. Prepare 3-hydroxypicolinic acid (3-HPA, 40 mg/mL) in mixture of acetonitrile/water (7:3, v/v). 3. Ensure complete dissolution by mixing the solutions well, sonication (5 min), and centrifugation. 4. Mix 90 µL of 6-HPA solution with 10 µL of 3-HPA solution and add 5 mg cation exchange resin (NH 4 ⫹ form). 5. Centrifuge, separate supernatant, and add 5 mg cation exchange resin to supernatant solution (MALDI matrix solution). 6. Transfer 0.5 µL collected sugar onto MALDI target and evaporate the sheath liquid for 5 min at 45°C. 7. Add 0.5 µL MALDI matrix solution with cation exchange resin to dried samples. 8. Operate MALDI/TOF instrument in negative mode (⫺18 kV) according to instrumental instructions

NOTES 1. The minimum flow rate necessary to prevent sample ions from migrating from the end of the capillary to the electrode, V min , can be calculated by following equation: V min ⫽ πµEBGEκBGEd 2 /4κCB, if µ is the mobility of the species, EBGE is the electric field strength inside the separation capillary, κBGE and κCB are the conductivities of the buffer inside the collection capillary and in the sheath flow tip and d is the ID of the separation capillary [see also Ref. 5]. Even flow rates of 1 µL/min will prevent fast biomolecules from migrating to the electrode. 2. The bands in CE contain usually only a few nanoliters of sample. Therefore, only a small sample amount can be collected during CE separations. 3. The elution time, te , can be calculated according to the following equation: te ⫽ (X/leff ) ⋅ tdet (X ⫽ distance between detector and capillary end, leff ⫽ effective capillary length, tdet ⫽ migration time between injection and detection point). The absolute error of the prediction of the elution time, ∆te , is proportional to the elution time. Therefore, the shorter the distance between detection point and capillary end, the smaller the error in predicting the elution time. 4. If collected samples are to be analyzed by mass spectrometry (i.e., MALDI/TOF), use of a volatile buffer is recommended to avoid formation of salts with MALDI matrix (organic acids). In general, all buffer solutions should be filtered for CE separations (0.2–0.45 µm pore size).

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Mu¨ller and Suzuki If possible the sheath buffer should be the same as the separation buffer (exception: cIEF runs). Desalted DNA samples can be injected electrokinetically using the stacking effect. This measure concentrates the sample during injection, thus increasing the amount of sample that can be loaded onto the separation capillary and minimizes injection dispersion. For purity check only short capillaries are required. On commercially available instruments short end injections are recommended. Attention! EtBr is a mutagenic substance. Handle with extreme care (gloves, etc.). EtBr should be added to the separation matrix and the anodic buffer vial. Staining of the dsDNA bands (intercalation) occurs during electrophoresis. Store at ⫺20°C when not in use. Sheath flow collection shows a special phenomenon which can not be observed in any other collection technique: backflow. This phenomenon arises from the fact that both ends of the separation capillary are immersed in different environments. One end is inside a small droplet, whereas the other end is inside a buffer reservoir (large volume). The surface tension of the small droplet creates a pressure inside the droplet, which is sufficient to create a flow in direction to the inlet buffer vial. This hydrodynamic flow can cause band broadening and has to be considered whenever working with a sheath flow cell. The band broadening expressed in height of a theoretical plate, H, is: H ⫽ r 2νbf /24D, where r is the radius of the separation capillary, νbf is the backflow velocity and D the diffusion coefficient of the species. Again, cIEF separations and separations in viscous polymer matrices are favorable. During cIEF separations band broadening is minimized by the constant refocusing of the zones. In viscous polymer matrices no band broadening can be detected, since the hydrodynamic flow is almost completely suppressed. The capillary should end flush with the Teflon tip as shown in Figure 1. It is impossible for the backflow to occur, while the capillary inlet is in the air (on the same level as the Teflon tip). The reason for this stems from the fact that if the buffer would flow out on the inlet end, a droplet would have to be formed. This droplet would in the first instance be smaller than the sheath droplet and therefore create a higher pressure directed in the opposite direction. Therefore the fluorescein solution stays inside the separation capillary until the capillary is inserted into the buffer vial. From the beginning of the run there will be first a plateau in the back-

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ground fluorescence. Then, after a while, the nonabsorbing sheath buffer reaches the detection window and decreases the background absorbance until it levels off in another plateau. The backflow velocity, νbf, can be calculated from: νbf ⫽ X/t1/2, with t1/2 being the time between the marked capillary insertion and the time when the background fluorescence decreased half the distance between maximum and minimum absorbance. The height of the inlet buffer vial does not have to be adjusted for separations in viscous polymer solutions. In all other cases it has to be remembered that the size of the sheath buffer droplet is changing constantly. Counterbalancing means that the overall flow velocity of the backflow is zero. This implies that there will be hydrodynamic flow in both directions, causing band broadening in CZE separations. To counterbalance back pressure and hydrostatic pressure, the capillary inlet has to be lifted a distance h, with h ⫽ 2σ/rgρ, where g is the acceleration by gravity, ρ is the density of the buffer, and σ is the surface tension of the sheath buffer. Since dissolution and degasing of methyl cellulose is a time-consuming process, it should be done in higher quantities. Methyl cellulose solutions can be stored for several months at 4°C. Alternatively, small quantities can be degased by centrifugation under vacuum. Replacing the separation matrix will elute the MC matrix at the end of the capillary. The viscous polymer might plug the collection capillaries and therefore has to be removed. It may be useful to have the first collection capillary connected to a mild vacuum (as shown in Fig. 3) to remove excess buffer during the run, before the first bands elute. In some cases it might be necessary to chose lower voltages or higher sheath flow rates in order to collect sufficient amounts of liquid. The collection capillaries will contain a collected band and waste in an alternating manner. It has to be noted that the sheath buffer will dilute the polymer matrix at the capillary tip. All bands will therefore elute several seconds too early. The collection software has to be set to an appropriate offset amount (calculated elution time minus 2–4 sec). Purity assessment is important especially for DNA applications, as impurities can greatly disturb PCR reactions or DNA sequencing. LIF detection is necessary to detect the collected fragments without further preconcentration, which have been diluted 500- to 1000-fold during collection. Lifting the inlet buffer vial will ensure that the ampholyte sample mixture is not pushed out of the capillary by backflow.

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20. The flow velocity can be calculated in analogy to Note 11 by ν ⫽ leff /t. A more accurate way of determining the mobilization flow velocity would be the use of two detection points close to the capillary outlet (online determination of flow velocity). 21. No offset value has to be considered in the software. 22. In the case of CZE separations there will be band broadening because of the parabolic velocity profile of the backflow (see also Note 13). It is recommended to perform an analytical reference run, showing optimal resolution. If the preparative run does not show satisfactory results a smaller capillary diameter can be chosen, which will decrease band broadening significantly. On the other hand, the amount of injected sample will decrease as well.

REFERENCES 1. Rose, D.J., Jorgenson, J.W. (1988) Fraction collector for capillary zone electrophoresis. J. Chromatogr. 438, 23–34. 2. Camilleri, P., Okafo, G.N., Southan, C., Brown, R. (1991) Analytical and micropreparative capillary electrophoresis of the peptides from calcitonin. Anal. Biochem. 198, 36–42. 3. Cheng, Y.-F., Fuchs, M., Andrews, D., Carson, W. (1992) Membrane fraction collection for capillary electrophoresis. J. Chromatogr. 608, 109–116. 4. Huang, X., Zare, R.N. (1990) Continuous sample collection in capillary zone electrophoresis by coupling the outlet of a capillary to a moving surface. J. Chromatogr. 516, 185–189. 5. Mu¨ller, O., Foret, F., Karger, B.L. (1995) Design of a high-precision fraction collector for capillary electrophoresis. Anal. Chem. 67, 17, 2974– 2980. 6. Guttman, A., Pritchett, T. (1995) Capillary gel electrophoresis separation of high-mannose type oligosaccharides derivatized by 1-aminopyrene3,6,8-trisulfonic acid. Electrophoresis 16, 1906–1911.

10 Active Microelectronic Array Systems for DNA Hybridization, Genotyping, Pharmacogenomic, and Nanofabrication Applications Michael J. Heller Nanogen, Inc., and University of California, San Diego, San Diego, California

Eugene Tu, Robert Martinsons, Richard R. Anderson, Christian Gurtner, Anita H. Forster, and Ron Sosnowski Nanogen, Inc., San Diego, California

10.1 INTRODUCTION A variety of miniaturized DNA arrays, DNA chips, Lab-on-a-Chip devices, and automated high-throughput screening (HTS) represent new technologies that will improve the way many important molecular biological analyses are performed in both research and clinical diagnostic laboratories. The development of these new technologies represents a synergistic combination of many disciplines that include microfabrication, microfluidics, MEMS, organic chemistry, molecular biology, genetics, and genomics. In some cases these new devices and technologies utilize sophisticated microfabrication processes developed by the semiconductor industry. Biochip arrays with large numbers of DNA test sites have been developed which utilize photolithography combinatorial synthesis techniques [1–3] or physical deposition methods to produce high-density DNA arrays [4,5]. Sequencing by hybridization in 223

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microarray formats [6–8] has been developed by other groups. Many of these DNA array technologies involve hybridization that occur simultaneously on the probe sequences attached to the surface of the array. In most cases, these DNA hybridizations are carried out under conditions where the reaction rates and stringency conditions are controlled by target DNA concentration, temperature, and salt concentration of the hybridization and wash solutions. These more classical or passive hybridization approaches limit the degree to which hybridization speed and selectivity can be improved. In many cases these passive array technologies may not meet the high performance hybridization criteria needed for carrying out some of the more challenging genotyping and gene expression applications. The development of microelectronic chip or array devices has overcome many of the limitations of passive hybridization techniques. These active electronic array devices facilitate: (1) the ability to produce reconfigurable electric fields on the microarray surface that allows the rapid and controlled transport of charged molecules to any test site; (2) the ability to carry out site selective DNA probe/target addressing and hybridization; (3) the ability to greatly increase DNA hybridization rate by concentration of probe/target DNA at the test site; and (4) the ability to use electronic stringency to improve hybridization specificity for point mutation, SNP, and STR analysis. In addition to DNA and RNA molecules, these devices have the ability to selectively transport any charged entity, which can include proteins (antibodies, enzymes, etc.), cells, nanoparticles, and semiconductor microstructures. These chips or arrays are microelectronic devices that exploit both microfabrication and microelectronic technology. More importantly, they are referred to as ‘‘active’’ electronic devices because they use electric fields—in particular, electrophoretic fields—to selectively transport DNA or other molecules and to directly affect the hybridization reactions or other affinity reactions occurring on the surface of the array [9–12]. 10.2 ACTIVE MICROELECTRONIC CHIPS AND ARRAYS Active microelectronic arrays have been designed and fabricated with 25, 100, 400, 1200, 1600, and 10,000 test sites or microlocations. The

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100-test site chip, which has been commercialized, has 80-µm diameter test site/microlocations with underlying platinum microelectrodes, and 20 auxiliary outer microelectrodes (see Fig. 1). The outer group of microelectrodes provides encompassing electric fields for concentrating DNA from the bulk sample solution to specific test sites/microlocations. Each microelectrode has an individual wire interconnect through which current and voltage are applied and regulated. The 100-test site DNA chip is about 7 mm in size, with the active test site array area being about 2 mm in size. Figure 2 shows a cross section through a typical 100-test site chip. The chips are generally fabricated on silicon wafers. The base structure is silicon with an insulating layer of silicon dioxide. The microelectrode structures are platinum and the connecting wires are gold. Silicon dioxide or silicon nitride is used to cover and insulate the conducting wires, but not the surface of the platinum micro-

Figure 1 An active microelectronic DNA array device with 100 test sites or microlocations. The test sites are ⬃80 µm in diameter, with an underlying platinum electrode. A ring of another 20 microelectrodes that can be used as counterelectrodes surrounds the test sites.

Figure 2 Cross section through the microelectronic array device showing the silicon base, the insulating silicon dioxide layers, the platinum microelectrodes, and the overlying permeation layer structure.

Figure 3 View of the automated spin coater system that is used in the microarray manufacturing process. This system carries out the controlled application of the permeation layer onto the microarray surface.

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electrode structures. The whole active array surface is covered with several microns of hydrogel (usually agarose or polyacrylamide) which forms the permeation layer. Usually the permeation layer is impregnated with a coupling agent (streptavidin, chemical agent, etc.) which allows for the subsequent attachment of DNA probes or other entities [13]. Figure 3 shows the automated system that is used to apply the permeation layer. This system allows the permeation layer to be applied rapidly and in a highly controlled manner. Ability to properly manufacture this key structure is important to the performance of the device for electronic hybridization. The ability to use silicon and microlithography for microfabrication of the DNA chips allows a wide variety of devices to be designed and tested. Figure 4 shows some of the unique DNA chips that have been designed and tested at Nanogen. In many cases the designs were used to test different microelectrode configurations as well as different manufacturing processes [14]. Figure 5 shows a silicon wafer of a 400-test site device with 50-µm microlocations and Figure 6 shows a 10,000-test site device with 30-µm microlocations. These represent more sophisticated devices that have on-chip CMOS

Figure 4 Other microelectronic arrays that have been designed and used to test various microelectrode configurations and microfabrication parameters.

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Figure 5 Wafer containing 400 test site active microelectronic array devices. These devices contain underlying CMOS control elements for on-chip current and voltage control and regulation.

Figure 6 View of an individual 10,000 test site active microelectronic array device. These devices contain underlying CMOS control elements for on-chip current and voltage control and regulation. The device has 30-µm test sites and is ⬃12 mm in size.

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control elements. In these devices, the current and voltages to the microelectrode at each test site are controlled and regulated by semiconductor elements on the chip itself [15]. These control elements are located in the underlying silicon structure and are not exposed to the aqueous samples that are applied to the chip surface when carrying out the DNA hybridization or other affinity reactions. 10.3 CHIP/CARTRIDGE AND ELECTRONIC HYBRIDIZATION SYSTEM The 100-test site microelectronic chip or array device is incorporated into a cartridge package (NanoChip cartridge) which provides for the electronic, optical, and fluidic interfacing. The NanoChip cartridge assembly is shown in Figure 7. The chip itself is mounted (flip chip bonded) onto a ceramic plate and pinned out for the electrical connections. The chip/ceramic plate component itself is mounted into a plastic cartridge that provides several fluidic input and output ports for addition and removal of DNA samples and reagents (fluorescent probes, buffers, etc.). The area over the active test site portion of the array is an enclosed sample chamber (⬃11 µL volume) covered with a quartz glass window. This window allows for fluorescent detection to be car-

Figure 7 Several views of the NanoChip cartridge device. The microelectronic array is incorporated into the cartridge package that provides the electronic, optical, and fluidic interfacing.

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ried out on the hybridization reactions that occur at the test sites on the array surface. A complete instrument system (NanoChip Molecular Biology Workstation) provides a chip loader component, fluorescent detection/reader component, computer control interface, and data display screen component (see Fig. 8). The probe-loading component allows DNA probes or target molecules (DNA, RNA, PCR, or other amplicons, etc.) to be selectively addressed to the array test sites, providing the end user with ‘‘make your own chip’’ capabilities. With the automated probe loader system four 100-test site NanoChip cartridges can be loaded with DNA probes and DNA samples from a 96- or 384-well microtiter plate (see Fig. 9). The probe loader component allows oligonucleotide probes, PCR amplicons, and DNA or RNA samples to be electronically addressed to the array in almost any desired arrangement. Probes or target sequences are usually in a biotinylated form, as this allows them to become attached (bound) to streptavidin within the permeation layer

Figure 8 Research system or NanoChip Molecular Biology Workstation. This system provides a chip loader component, fluorescent detection/reader component, computer control interface, and data display component.

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Figure 9 Probe loader component used for automated DNA probe and DNA target sequence addressing or spotting. This component provides the end user with ‘‘make your own chip’’ capabilities.

at the specified test site. By way of example, Figure 10 shows how a group of 25 test sites on a microelectronic array could be loaded (addressed) with five different biotinylated oligonucleotide probes, at each of five specific test sites in five different columns. In the electronic addressing procedure the probe loader component automatically delivers the desired biotinylated oligonucleotide probe to the array, biases the specified test sites positive and counterelectrodes negative. The electric (electrophoretic) field causes the biotinylated probes to concentrate onto the positively activated test sites, with subsequent binding via the biotin/streptavidin reaction [13,16]. Nonspecific binding of the biotinylated probes to any nonbiased or negatively biased test sites does not occur under the electronic addressing conditions. The procedure is repeated for all five probe sequences until the array is completely addressed. The probe loader device allows the end user to essentially address the array in any desired fashion. Electronic addressing also

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Figure 10 Scheme for selective electronic addressing (spotting) of biotinylated oligonucleotide probes or target DNA sequences to specified test sites on the array.

allows DNA probes to be spotted onto the array in a highly reproducible manner. Figure 11 shows an example of the precision achieved by electronic probe addressing. In this case, a fluorescent biotinylated probe was repeatedly electronically addressed 100 times with a resulting CV of ⬃5%. This type of reproducibility would be hard to achieve with more conventional probe spotting techniques. Microelectronic arrays can be formated in a variety of ways that include reverse dot blot format (capture/identity sequences bound to test sites), and dot blot format (target sequences bound to test sites). After an array has been addressed (capture probe or target DNA loaded), various electronic hybridization assays can be carried out and the chip subsequently analyzed using the fluorescent detection system. Figure 12 shows an example of an editing map for probe loading and electronic hybridization. The left side (Fig. 12) shows the map for one possible capture probe loading scenario, with different colors used to represent the oligonucleotide capture probes in the 96-well microtiter

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Figure 11 An example of the precision achieved by electronic probe addressing. A fluorescent biotinylated probe was repeatedly electronically addressed 100 times with a resulting CV of ⬃5%.

plate and their specified test site position on the microelectronic array. The right side (Fig. 12) shows the editing map for selective hybridization of target or sample DNA to specific test sites on the array. The different colors are used to represent the different target or sample DNAs in the 96-well microtiter plate and their specified positions for electronic hybridization on the array. The lower figure shows the combined addressing and sample loading map, which allows the end user to keep track of even the most complicated addressing and hybridization procedures being carried out on the array. Hybridization assays carried out on the research system involve the use of fluorescent reporter probes or target DNA sequences. The reporter groups are usually organic fluorophores that have been attached either to oligonucleotide probes or to the target/sample DNA/ RNA sequences. After electronic addressing and hybridization are carried out, the chip is analyzed using the fluorescent detection system. The fluorescent detector has two different laser excitation sources (ex-

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Figure 12 Probe loader and electronic hybridization editing maps. Appropriate software and data display keeps track of where probes have been electronically addressed to the array, and where sample or target DNAs are to be hybridized.

citation 532 nm and excitation 635 nm). The laser beams are quickly scanned across the array using a confocal type optical system and the emissions from the fluorescent labeled probes or targets (550–600 nm ‘‘green’’ and 660–720 nm ‘‘red’’) are detected. Figure 13 provides an example of adrenoreceptor SNP genotyping analysis, and shows what the fluorescent image of the hybridized array looks like. In this example, test samples were first selectively addressed to the chip (dot blot/ sample down format), and then the array was hybridized with red fluorescent (mismatch) and green fluorescent (match) reporter probes. The left-hand view (Fig. 13) shows a fluorescent image of the array using the green detection channel, and the right-side shows the fluorescent image for the red detection channel. The lower graph (Fig. 13) shows the fluorescent data analysis results for the adrenoreceptor

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Figure 13 Fluorescent analysis and data presentation for Adrenoreceptor SNP genotyping (sample/target down format) using 100 test site microelectronic array (NanoChip cartridge) and research system (NanoChip Molecular Biology Workstation).

SNP genotyping of each test sample. A predominant green fluorescent signal (bar) with minimum red signal represents a homozygous wildtype allele (normal) call for the genotype. A predominant red signal (bar) with minimum green signal represents a homozygous mutant allele call for the genotype. A nearly equivalent green and red signal (bars) means a heterozygous call for the genotype. Genotyping analysis using electronic hybridization and stringency provides very high discrimination ratios and unambiguous calls. Selection rules are simple, and no complex image processing is required for data analysis. Genotyping calls are easy to make and results are highly reliable.

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10.4 ASPECTS OF ELECTRONIC HYBRIDIZATION Selective electronic addressing of DNA probes and subsequent electronic hybridization reactions on the active microelectronic array is carried out by application of a positive DC bias to the individual microelectrodes beneath the selected test sites while maintaining other selected countermicroelectrodes negative. These include the outer group of 20 countermicroelectrodes that surround the actual test site microlocations. The application of the appropriate current and voltage levels to the underlying microelectrodes produces an electrophoretic field, which affects the transport and concentration of the negatively charged nucleic acid molecules to the selected (positively biased) test sites on the array. The microelectronic array is designed so that the test sites or microlocations on its surface can be negatively or positively biased in any desired arrangement. Thus, almost any pattern of DNA movement to one or more test sites on the array can be carried out. Functionalized nucleic acid probes (oligonucleotides, DNA, RNA, PCR amplicons, polynucleotides, etc.) can be selectively and rapidly immobilized and bound (covalently or noncovalently) within the permeation layer of the microlocation overlaying the activated platinum microelectrode. Similarly, the target nucleic acid sequences in the sample can then be transported, concentrated, and hybridized to the desired test-sites previously addressed with the nucleic acid probes. This rapid concentration of target DNA at the microscopic test sites leads to a significant reduction in the hybridization time when compared to passive hybridization techniques. For many assays where the concentration of target DNA in the sample is low, hybridization reactions occur in seconds rather than hours. Reversal of the electric field potential (negative bias) at the test site will now cause the rapid removal of unhybridized DNA molecules. When the electric field is adjusted to the appropriate level, it can be used to affect the selective dehybridization of the hybridized DNA sequences from the attached complementary probe. This novel parameter is called ‘‘electronic stringency’’ and it provides a powerful and rapid method for single base mismatch discrimination analysis of point mutations and SNPs [13,16,17].

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Facilitating the electrophoretic transport, hybridization and stringency processes on the active microarray depends on several important techniques and device features. First, a key role is played by the permeation layer, which overlies the microelectrodes. The permeation layer is of critical importance in the case of planar microelectronic array devices where the microelectrode surface is in close proximity to the DNA capture probes and hybridization reactions [13,16]. Generally, the permeation layer is a thin coating of a hydrogel (agarose or polyacrylamide) which is spin-coated over the chip surface. The permeation layer is usually ⬃1–2 µm in thickness in the dry state, and swells to ⬃5 µm when hydrated. The permeation layer allows the array to be run at current levels (⬎100 namp) and voltages (⬎1.2 V) which provide electrophoretic transport, while serving to protect the sensitive DNA hybridization reactions from the adverse electrochemical effects which occur on the actual microelectrode (platinum) surface during active operation. These adverse effects include the generation of hydrogen ions (H ⫹) and oxygen at the positive microelectrode surface and hydroxyl ions (OH ⫺) and hydrogen at the negative microelectrode surface, as well as various free radical entities that are created by the electrolysis process [13,16]. The permeation layer also serves as a matrix for the attachment of DNA capture probes and target DNA sequences. Attachment is achieved by impregnating the permeation layer with an affinitybinding protein like avidin or streptavidin for subsequent attachment of biotinylated probes, or chemically modifying the layer for subsequent covalent attachment of activated or modified DNA probes. Generally ⬃10 9 probes or target DNA (amplicons, RNA, etc.) sequences can be bound to the permeation layer at the 80-µm test site/microlocation (see Fig. 2). The structure of typical arrays and permeation layers has been described previously [9–11,13,16,17]. Another important consideration for active electronic transport and hybridization is the buffer composition. To facilitate both the rapid electrophoretic transport of DNA molecules and their efficient hybridization, special zwitterionic buffer species with low-conductivity properties have been utilized. While many low-conductance zwitterionic buffers can be used to promote the rapid transport of charged molecules (DNA) on the array, most cannot generate sufficient cationic property

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to stabilize the hybridization process. Histidine has been found to be particularly effective for electronic hybridization. Histidine in its zwitterionic form at or near neutral pH has a very low conductivity, provides relatively good buffering capacity, and also stabilizes hybridization when protonated. Histidine at 50 mM concentration, in its zwitterionic state typically has a conductivity of ⬍100 µS/ cm, while conventional buffers commonly employed in hybridization analysis (6 ⫻ sodium chloride/sodium citrate) have conductivities 1000-fold greater [11]. Figure 14 shows the proposed mechanism for electronic hybridization using zwitterionic histidine buffer. Stabilization of hybridization at the test site is believed to be due to the buffering

Figure 14 Mechanism for electronic-based hybridization in zwitterionic Histidine buffer. The figure shows a cross section of a test site with the platinum microelectrode and permeation layer with immobilized capture probes. During the application of a positive bias to the microelectrode, the resulting electrophoretic field begins to concentrate the target DNA in the bulk solution at the activated test site. The buffering effect of zwitterion Histidine at the positive biased microelectrode produces cationic Histidine molecules which then stabilize the hybridization of the target DNA sequences to the bound DNA probes at the activated test site.

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effect of zwitterionic histidine at the positive biased microelectrode. This buffering effect produces dicationic histidine which then apparently stabilizes the hybridization of the target DNA sequences which have been concentrated near the bound DNA probes at the activated test site. Electronic hybridization in zwitterionic histidine buffer also provides several other advantages, which include: (1) target DNA in the sample solution remains substantially denatured; (2) competitive hybridization from complementary strands is reduced; and (3) secondary DNA/RNA structure is broken. Table 1 provides a list that summarizes many of the advantages of electronic hybridization. These advantages have proven particularly important for carrying out genotyping of problematic point mutations, SNPs, and STRs. Microelectronic arrays and electronic hybridization techniques can be used for a number of important research, molecular diagnostic, and pharmacogenomic applications. These include a variety of genotyping applications such as point mutation, single-nucleotide polymorphism (SNP), and short tandem repeat (STR) analysis. In almost all cases electronic hybridization provides highly accurate and reliable results. In particular, electronic hybridization allows more reliable results to be achieved for problematic SNPs that are difficult to genotype by conventional array hybridization techniques. Other, nearer-term applications which can be carried out on microelectronic arrays include gene expression and on-chip amplification. Microelectronic arrays may also Table 1 Advantages of Electronic-Based Probe and Target Addressing and Hybridization • Rapid probe addressing, hybridization, and stringency • Site selective addressing, hybridization, stringency at any time in experiment • Electronic addressing and hybridization occurs only at active test sites; ‘‘no passive addressing or hybridization occurs at any other position on the array’’ • Electronic hybridization with low conductance buffers advantages: (1) DNA substantially denatured; (2) little competition from complementary strands (PCR amplicons); and (3) secondary DNA/RNA structure is broken

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Table 2 Various Electronic Techniques for DNA Hybridization, Molecular Diagnostic, and Pharmacogenomic Applications (1) Single-nucleotide polymorphisms (SNP), rapid with high accuracy and reliability (2) Short tandem repeats (STR) for forensic, ID, and diagnostics (microvariants) (3) Gene expression (4) Combined genotype (SNP/STR) (5) Combined genotype/gene expression and/or phenotype (6) On-chip in situ amplification (7) Cell separation for research and POC systems

be useful for combining different molecular diagnostic assays together, for example: SNPs and STRs; genotyping and gene expression; and genotyping, gene expression, and phenotyping. Still other applications include immunodiagnostics, mutation scanning, cell separation, and the development of sample to answer processes for point of care diagnostic systems (see Table 2). 10.5 GENOTYPING APPLICATIONS 10.5.1

Single Nucleotide Polymorphisms

Electronic hybridization allows most genotyping applications, which can include point mutations, single nucleotide polymorphisms (SNPs), and short tandem repeats (STRs) to be carried out rapidly with high accuracy and reliability. Genotyping can be carried out in a variety of hybridization formats which include reverse dot blot, sandwich and sample, or target down (dot blot). In the reverse dot blot format, a set of capture probes that are complementary to the match and mismatch bases in the target sequence are selectively addressed and bound to the

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array. The capture probes are usually biotinylated synthetic oligonucleotides from 15 to 25 nucleotides in length. Labeled target DNA is then electronically hybridized to the specific capture probes on the array. Target DNA can be basically any size, with sequences up to 1000 nucleotides having been genotyped. In the case of target or sample DNA from a PCR reaction, a fluorescent label can be incorporated via use of a PCR primer that has been derivatized with a suitable fluorescent moiety (Bodipy Texas Red, Cy5, etc.). Alternatively, the hybridized target sequence can be secondarily hybridized with a fluorescent-labeled reporter probe. After hybridization, various stringency techniques can be used to distinguish the match from the mismatch (discriminate the mutation); these include electric field stringency, temperature, ionic strength, pH, and extensive washing in various buffer solutions. The distinguishing characteristic of the reverse dot bot is that the match/mismatch specificity (discrimination) is in the capture probe sequences. In the sandwich format, capture probes that are specific for a portion of the target sequence are selectively addressed and bound to the array. Target DNA (PCR amplicon, etc.) is then hybridized to the array. The ‘‘captured’’ target DNA is then hybridized with a set of fluorescent reporter probes that are specific for the match and the mismatch bases in the target DNA sequence. In the case of base stacking type assays, the captured target DNA is hybridized with a stabilizer or helper probe and then with a set of fluorescent reporter probes [18]. Various stringency techniques are then used to distinguish the match from the mismatch (discriminate the mutation); these include electric field stringency, temperature, ionic strength, pH, and extensive washing in various buffer solutions. The distinguishing characteristic of the sandwich assay format is that the match/mismatch specificity (discrimination) is in the reporter probe sequences. For the base stacking-type assay, the specificity is in both the helper and reporter probes. In the target or sample down format (dot blot), the target or sample DNA (PCR amplicon, etc.) is first selectively addressed and bound to the array. The target or sample DNA is then hybridized with a set of fluorescent reporter probes that are specific for the match and the mismatch bases in the target sequence. In the case of base stacking-type assays, the target DNA is hybridized with a stabilizer or helper probe

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and then with a set of fluorescent reporter probes [18]. Various stringency techniques can then be used to distinguish the match from the mismatch (discriminate the mutation); these include electric field stringency, temperature, ionic strength, pH, and extensive washing in various buffer solutions. The distinguishing characteristic of the target or sample down format is that the match/mismatch specificity (discrimination) is in the reporter probe sequences. For the base stacking-type assay, the specificity is in both the helper and reporter probes. An example of a target or sample down (dot blot) hybridization assay for Hemochromatosis is shown in Figure 15. In this case, PCR amplification was carried out for the Hemochromatosis gene segment in 16 different test samples. One of the PCR primers in the set is biotinylated, and this amplicon strand is the actual target sequence that becomes bound to the array test site. The PCR samples were then electronically addressed to the selected test sites on the array. The array

Figure 15 Example of the target or sample down (dot blot) electronic format for hybridization and discrimination of point mutations for Hemochromomatosis in 16 different samples.

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was then hybridized with a set of fluorescent reporter probes for the different Hemochromatosis alleles (see sequences in Fig. 15). The results show that samples 3, 4, 8, 9, 11, 13, and 16 are the homozygous wild-type allele (normal) for the Hemochromatosis genotype; samples 2, 5, 6, 10, 14, and 15 are homozygous mutant allele for Hemochromatosis genotype; and samples 1, 7, and 12 are heterozygous. Because of the high discrimination ratios, all the sample calls are easy to make and unambiguous. Figure 16 shows another example of target or sample down (dot blot) hybridization, in this case for multiplexed Hemochromatosis and Factor V genotyping. Multiplex PCR amplification was carried out for Hemochromatosis and Factor V in three different samples. One of the primers in each of the primer sets for Hemochromatosis and Factor V is biotinylated. The three multiplexed PCR samples were then electronically addressed to the array. The array was then hybridized with a set of fluorescent reporter probes for the Hemochromatosis alleles and then with a set of fluorescent reporter probes for the Factor V alleles. The results show that the first sample is a homozygous wild-type allele for Hemochromatosis and homozygous mutant allele for Factor V, the second sample is heterozygous for both Hemochromatosis and Factor V,

Figure 16 Results for multiplex electronic hybridization and point mutation analysis for Hemochromatosis and Factor V in three different samples.

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and the third sample is homozygous wild-type allele for Hemochromatosis and homozygous wild-type allele for Factor V. Again, the calls are unambigouous because of the high discrimination ratios. A variety of stringency techniques can then be used to discriminate point mutations or other types of mutations (insertions, deletions, etc.) when doing genotyping analysis. This includes electric field stringency which is unique to the electronic array technology and systems, as well as more classical parameters of temperature, ionic strength, pH, and extensive washing in various buffer solutions. Electronic stringency involves using the electric field with reverse bias (negative) to remove unbound DNA and in the case of point mutations and SNPs to remove the hybridized mismatched probes relative to the hybridized matched probe. Gilles and other workers have demonstrated the advantages of electronic stringency for genotyping human mannose-binding (MBP) protein single-nucleotide polymorphisms [17]. In this work the electronic array based genotyping results for 22 blinded MBP quadraallelic samples and 13 blinded D allele samples were in 100% agreement with conventional DNA sequencing results. 10.5.2

Short Tandem Repeats

Short tandem repeats (STRs) represent another type of polymorphism with important applications in forensics, human identification, and diagnostics. In the case of forensics a standard set of polymorphic loci have been chosen for human identification. The criterion for the selected loci is their relatively high level of heterozygosity, which allows the generation of statistically unique ‘‘DNA fingerprints’’ with a minimum number of loci. The typical STR loci are selected groups of four nucleotide repeats that are represented in the human population by four to 15 alleles [19,20]. These alleles are distinguished by a different number of repeat units that may also contain point mutations. Unlike singlenucleotide polymorphisms (SNPs), STRs are more difficult to analyze by hybridization techniques and are generally not done using microarray technologies that rely on passive techniques [21,22]. However, electronic hybridization techniques have been proven to overcome these problems and allow STR analysis to be reliably performed on microelectronic arrays [18].

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Figure 17 Electronic STR genotyping analysis showing the homozygous THO 9.3 allele, the heterozygous TPOX 8/9 allele, and the heterozygous CSF 9/10 alleles.

An example of STR analysis using electronic hybridization on a microarray is shown in Figure 17. This shows the results for one sample (individual) which was tested for three STR loci that included THO (CATT), TPOX (AATG), and CSF (AGAT). The STR analysis of the particular sample (individual) shows that they are homozygous for the THO 9.3 allele (which includes a point mutation), heterozygous for TPOX with eight and nine allele repeat units, and heterozygous for CSF with nine and 10 allele repeat units. These results demonstrate the ability of electronic hybridization on microarrays not only to effectively resolve short tandem repeats, but also to resolve the microvariants (point mutations) within the repeats. Thus, microelectronic arrays can provide a useful tool for carrying out rapid and highly reliable forensic and human identification applications. 10.5.3

Genotyping Accuracy and Reliability

In addition to speed, genotyping by electronic hybridization can also provide accurate and highly reliable results. These particular perfor-

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mance criteria will be extremely important for future clinical diagnostic applications, where the generation of a false-positive and/or falsenegative result is unacceptable. Genotyping errors can occur frequently when using conventional techniques such as passive hybridization arrays, RFLP analysis, and DNA sequencing. It is more problematic that these errors are often not discovered until samples are reanalyzed a number of times or corroborated by other technologies. Figure 18 shows the electronic hybridization results for the SNP analysis of 32 samples for the Diastrophic Dysplasia (DTD) sulfate transporter polymorphism (Dr. Glen Evans, private communication). Each nucleotide for the biallelic polymorphism is graphed using a different color (T is red, C is green), which represents the normalized fluorescent intensity signals measured at the specific test sites on the microarray. Results for the DTD genotyping analysis carried out on a microelectronic array were 100% accurate. By way of example, samples 8, 9, 10, 11, 13, 14,

Figure 18 Results for DTD SNP analysis of 32 patient samples done by electronic hybridization and compared to the DNA sequence analysis results. No call (NC) and discordant calls (*) are marked. Electronic hybridization proved 100% accurate.

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18, 19, 21, 23, 24, 29, 31, 32, 34, 35, 36, and 38 are homozygous wildtype allele calls, having primarily green signal (bars) with very little red signal. The two samples 16 and 37 are homozygous mutant allele calls having primarily red signal with very little green signal. Samples 7, 17, 15, 20, 22, 25, 26, 27, 28, 30, and 33 are heterozygous calls, having nearly equal green and red signals. Sample 12, with a disproportionate red and green signal, was given a ‘‘no call’’ status by the electronic hybridization discrimination criteria; this sample was later determined to have been a mixed sample. Several of the initial sequencing calls, samples 24* (c/t), 28* (t/t), and 36* (c/t) were initially discordant with the electronic hybridization calls 24 (c/c), 28 (c/t), and 36 (c/c). After resequencing, all of the discordant calls were resolved in favor of the electronic hybridization calls. Thus, for this group of samples, the electronic hybridization technique produced results with higher accuracy and reliability than those produced by DNA sequencing, which would have been considered the gold standard technology. In general, electronic hybridization on microarrays may produce more accurate results for genotyping than conventional arrays and/or other techniques (sequencing, RFLP, etc.). This may come from the fact that electronic hybridization seems to be particularly advantageous for analyzing difficult or problematic polymorphisms. The case is clear for STRs where there is little data showing that this type of polymorphism can be analyzed by conventional passive array hybridization, and where electronic hybridization produces excellent results including the ability to type the microvariants within the repeats [18]. Regarding point mutations, SNPs, insertions, and deletions, electronic hybridization may provide advantages by overcoming some of the more intrinsic problems of the hybridization process itself. For example, in some cases a mutation may be associated with a region of a DNA sequence that is itself more difficult to hybridize. Figure 19 shows an example of one type of problematic SNP where there is a clear rationale for the advantage of electronic hybridization. This is an example of a SNP which is located near or in a region with strong secondary structure (stem loop). As shown in the upper part of Figure 19, a mutation near or in a region with strong secondary structure is relatively difficult to analyze by conventional hybridization techniques. Electronic hybridization which is carried out under substantially denaturing conditions

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Figure 19 Example of secondary structure (stem loop) problem that can prove difficult for conventional microarray analysis, but is easily overcome when using electronic hybridization.

is effective in eliminating the secondary structure problem and allows efficient and specific hybridization to take place (see lower section, Fig. 19). This ability to carry out hybridization under substantially denaturing conditions as well as a number of the other advantages of electronic hybridization is given in Table 1. Problematic or difficult genotyping may have a number of other causes, which can include mutations in high GC-containing sequences, mutations near or in repeat sequences, mutations located in close proximity, and mutations in large DNA fragments. In most of these cases electronic hybridization has been shown to produce accurate and highly reliable results. This type of performance criteria may be particularly important for carrying out clinically relevant molecular diagnostics. 10.5.4

Combined SNP and STR Genotyping

Different genotyping tests can be combined on a single microarray to provide enhanced flexibility of assay formats, which may also be important for future DNA diagnostics. In experiments done in conjunction with human identification projects, both SNPs and STRs were analyzed on the same chip. Figure 20 shows the results for combined SNP and

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Figure 20 Example of carrying out combined SNP and STR electronic hybridization analysis on the same chip. The first panel shows the array after hybridization, and the second and third panels show the array after stringency. The second panel is the image from the green detection channel, and the third panel is the image from the red detection channel.

STR genotyping on the same microarray. The first frame of Figure 20 (left side, upper section) shows the hybridization results for the STR loci CSF and TPOX before stringency has been applied. Because of the unique attributes of electronic hybridization, it was possible to develop a novel method of STR discrimination based solely on hybridization. The STR array is made up of all the alleles of the two loci (CSF and TPOX) with duplicate pads for each allele. Discrimination of the alleles occurs by taking advantage of base stacking within concordant hybridization complexes [18]. The lower part of the first frame in Figure 20 shows the SNP targets hybridized in duplicate for the EH1, Factor V, and Hemochromatosis genes. The top row of this section of the microarray has the unknown target; the bottom row has control DNA sequence. Two differently fluorescent reporter probes are used for each SNP. One reporter (green) will form a more stable hybrid with one allele (wild type), while the other reporter (red) will form a more stable hybrid with the other allele (mutant). Therefore, by analyzing the microarray with the two different fluorescent detector channels (red and green) after stringency has been applied, the genotype of the unknown target can be determined.

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The second two sections of Figure 20 now show the microarray after stringent conditions have been applied. The two panels in Figure 20 (center and right) are of the same array, but are the images from the red and green detector channels. While two-color detection is relevant to the SNPs on the bottom half of the array, the STR allele discrimination relies on the position of the signal rather than the color. Therefore, the STR analysis utilizes only the green channel. For the CSF alleles, stringent conditions have removed signal from all but four pads. The experiment was done with duplicate pads, so the result means that the unknown sample contained two CSF alleles, the 10 and 11 repeat units, and was therefore heterozygous. The TPOX sample shows only one signal at a single set of duplicate pads, meaning that the sample was homozygous for the eight repeat units. To determine the SNP genotype, the results from both the red and green channels must be used. The presence or absence of signal is determined relative to the heterozygote controls. The EH1 sample is identical to the heterozygote control in

Figure 21 Combined genotyping results for the STR analysis showing the sample was homozygous for TPOX 8 allele and heterozygous for CSF 10/11 alleles.

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both the red and green channels, indicating that it contains both alleles and is heterozygous. The Factor V sample has signal only in the green channel, indicating that it is homozygous wild type, and the Hemochromatosis sample has signal only in the red channel, indicating it is homozygous mutant. The fluorescent signals from STR test sites on the microarray that were shown in Figure 20 were quantified, and the results are shown as graphs in Figure 21. This figure shows the amount of signal remaining after stringency was applied to the test sites for each of the different STR alleles (repeats). For the TPOX locus there is the possibility for eight different four-base (AATG) repeat alleles present in the human population that are repeated from six to 13 times. For the CSF locus there is the possibility for nine different four-base (AGAT) repeat alleles present in the human population that are repeated from seven to 15 times. The X-axis on the graphs in Figure 21 indicates the number of STR repeat units potentially present for each test site. For the TPOX

Figure 22 Combined genotyping results for the SNP analysis showing the sample was homozygous mutant allele for Hemochromatosis, homozygous wild-type allele for Factor V, and heterozygous for EH-1.

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sample, the signal remains only at the eight repeat unit test sites, indicating that the sample was homozygous for that allele. For the CSF sample, the signal remains at both the 10 and 11 repeat unit test sites. Thus, this sample contained DNA sequences with two different CSF alleles and is heterozygous. Figure 22 is a graph that shows the results from the SNP samples. This graph shows that the sample DNA was a homozygous mutant allele for Hemochromatosis, a homozygous wild-type allele for Factor V and heterozygous for the EH-1. The ability to carry out combined SNP and STR genotyping analysis represents a unique formatting advantage of electronic hybridization on microarrays. This type of formating could be extremely useful for future forensic, human ID, and clinical diagnostic applications.

10.6 OTHER DIAGNOSTIC APPLICATIONS AND SYSTEMS 10.6.1

Immunoassays and Combined Assays

Active microelectronic array systems can be used for a wide variety of DNA hybridization, molecular biological, and clinical diagnostic applications. In addition to SNP and STR genotyping applications already described, other applications include gene expression analysis, on-chip or in situ DNA amplification [23,24], immunoassays [25,26], proteinprotein interactions, viral and bacterial identification, biowarfare agent detection, cell separations [12,26,27], pharmacogenomic and drug discovery applications, and micro/nanofabrication [28,29]. An example of carrying out an immunochemical assay in combination with a genotyping assay is shown in Figure 23. The upper portion of the fluorescent image shows a specific antibody-antigen immunoassay for alpha-bungarotoxin. The experiment was carried out by initially addressing and binding a biotinylated anti-alpha-bungarotoxin antibody (capture antibody) to two test sites on the array and a nonspecific antibody to two other test sites that serve as a negative control. The alpha-bungarotoxin antigen was then electronically addressed to both specific and nonspecific antibody test sites on the array. The capture/target immunocomplex was then addressed with a fluorescent

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Figure 23 Combined electronic-based immuno/DNA toxin assay for alphabungarotoxin and for Shiga-like toxin.

secondary antibody. The anti-alpha-bungarotoxin antibody-antigen complex was formed as is shown in the upper left corner of Figure 23. Also shown in Figure 23 is a DNA hybridization assay for a Shigalike toxin gene target. This combined immunoassay/DNA genotyping toxin assay was done on an active microelectronic array using all electronic conditions for both assays (transport, affinity binding, and hybridization). Another, even more complex electronic-based immunoassay is shown in Figure 24. In this case the microarray was first addressed with four different capture antibodies that included bungarotoxin, antirabbit, anti–hep B, and the anti-choleratoxin MAB. The array was then addressed with choleratoxin, the target antigen. Finally, the array was addressed with the fluorescent reporter antibody, which detects the specific choleratoxin/anticholeratoxin MAB complex. Thus, electronic techniques can be used to carry out even more complex immunoassays as well as combined assays that include genotyping. Combined genotype/immuno as well as other genotype/phenotype assays provide a great deal of flexibility and potential for designing a variety of very

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Figure 24 An electronic-based immunoassay for choleratoxin. The array has been addressed with four different capture antibodies.

unique assays. Such assays may prove particularly useful for clinical diagnostic applications. 10.6.2

Integrated Sample-to-Answer Systems

The continued evolution of ‘‘Lab on a Chip’’ systems is being driven by both advancements in MEMS, microlithrography, and other miniaturization technologies, as well as the needs of end users in forensics areas, clinics, and hospitals to have miniaturized devices and systems which can carry out complicated diagnostic analyses. Figure 25 shows a sequence of technology development going from bench-top manual assays to an automated system (Nanochip Molecular Biology Workstation), to miniaturized amplification systems, to sample-to-answer automated systems and eventually leading to a highly integrated labon-a-chip device. One of the major goals in this area is to develop miniaturized ‘‘sample-to-answer’’ and ‘‘point-of-care’’ systems which would have applications in doctor’s offices, emergency rooms, field diagnostics, environmental monitoring, and biological warfare agent monitoring. These applications as well as the desire to make assays more sensitive, more specific, faster, less expensive, smaller, and more

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Figure 25 Evolution of ‘‘Lab-on-a-Chip’’ technology shows the progression from completely manual assays and procedures carried out on the lab bench to a completely miniaturized and fully integrated lab-on-a-chip device.

automated are pushing the technology constantly toward more miniaturized and highly integrated systems. Figure 26 shows a prototype for a fully integrated sample-toanswer system. This miniaturized system is designed to first carry out cell separation (bacteria from blood), lyse the selected bacterial cells, amplify the extracted bacterial DNA, and then carry out the bacterial target DNA analysis on a microelectronic array [12]. Figure 26A shows the complete integrated system which is operated by a lap-top computer. Panel B shows the electronic cell separation/amplification array device (left side) and the microelectronic DNA hybridization array (right side). After cell separation (panel C) and lysis is complete, the amplification reagents are pumped into the first chip chamber which also contains a heating element. The bacterial target DNA amplification is allowed to take place. The amplified target DNA is then pumped out

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Figure 26 A prototype of an integrated sample to answer device that carries out cell separation (bacterial from blood), DNA amplification, and DNA hybridization analysis on a microelectronic array. (A) The whole device which is run by an associated lap-top computer;(B) the cell separation array (left side) and the DNA hybridization array (right side); (C) the separated bacteria cells (white) and blood cells (red); (D) the laser diode and CCD detector components.

of this chamber and into the microelectronic DNA array chamber which then carries out the electronic hybridization analysis using the appropriate fluorescent reporter probes. Fluorescent detection and analysis are now carried out by exciting the microarray test sites with a laser diode excitation source and monitoring the fluorescent signals with a CCD detector. The fluorescent signals are processed and a lap-top computer carries out data presentation. The laser diode and CCD detector components can be seen in panel D. A more complete description of

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this miniaturized integrated sample-to-answer system is described by Cheng and coworkers [12]. For certain applications the cell separation process can be a particularly important challenge for sample-to-answer systems. Many diagnostic applications involve trying to determine a very small amount of a specific analyte in a large background of extraneous material. Such examples include trying to identify a small number of specific bacteria or virus in a blood sample (infectious disease), fetal cells in maternal blood (genetic diseases), or tumor cells in a background of normal cells (early cancer detection). While a number of techniques are available, electronic-based methods have been developed that can have considerable advantages. One basic electronic method for cell separation is called dielectrophoresis. This process involves the application of an asymmetric alternating current (AC) electric field to the cell population [30,31]. Active microelectronic arrays have been used to achieve the separation of bacteria from whole blood [12] and for the separation of cervical carcinoma cells from blood [27]. Figure 27 shows the electronic separation of bacteria from whole

Figure 27 Separation of bacteria (E. coli) from whole blood. First panel shows array before application of sample. Second panel shows separation of bacteria and blood after AC electric field is applied. Third panel shows bacteria cells remaining at microlocations after blood cells have been washed away.

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blood. The left panel shows six test sites of a microelectronic array before introduction of a sample. The center panel shows the cell separation patterns which occur for a mixture of blood and bacteria after a high frequency AC electric field is applied in an asymmetric fashion. The upper and lower center panels show the different separation patterns that the blood cells take in the low field regions which is dependent on the asymmetry arrangement for the microelectrodes. In both cases, however, the bacteria are held in the high field region which is close to the microelectrode surface. While maintaining the AC field, the microarray is washed with a buffer solution that removes the blood cells (low field regions) from the more firmly bound bacteria (high field regions) near the microelectrodes. The bacteria can then be released and collected or electronically lysed to release the genomic DNA or RNA for further manipulation and analysis [12]. Dielectrophoresis represents a particularly useful process that allows difficult cell separation applications to be carried out rapidly and with high selectivity. This electronic-based process is also ideally suited for incorporation into integrated sample-to-answer systems and point-of-care systems.

10.7 ELECTRIC FIELD–ASSISTED MICRO/NANOFABRICATION 10.7.1

Enabling Nanotechnology

Concepts for nanotechnology encompass a wide variety of potential applications. Generally, molecular or nanoelectronic devices and systems are envisioned as the more revolutionary application of this new technology. Many examples of individual molecular components with appropriate basic properties now exist; these include entities such as carbon nanotubes and various organic molecules with electronic switching capabilities. The larger issue with enabling molecular electronics is more likely to be the development of a viable technology which will allow billions of molecular or nanoelectronic components to be assembled and interconnected into useful logic/memory devices and systems [32]. In addition to electronic applications, nanodevices and nanosystems with higher-order photonic, mechanical, mechanistic,

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sensory, chemical, catalytic, and therapeutic properties are also envisioned. Again, key problems with enabling such devices and systems will most likely occur at the stage of organizing components for higherlevel functioning, rather than the availability of the molecular components. By way of example, hundreds of relatively simple chromophore molecules are arranged in photosynthetic chloroplast structures which efficiently collect photonic energy and transfer it through these solidstate antennalike structures with high quantum efficiency. Figure 28 presents one of the challenges of nanofabrication by comparing a plant chloroplast with a CCD array—nature’s photonic device with a manmade photonic device. The plant chloroplast represents a highly integrated light-capturing device composed of numerous self-organized molecular, nano-scale, and micron-scale structures and components. The CCD represents a man-made photonic device in which the feature size stops at about the micron-scale level. In addition to the chloroplast being a true nano/molecular-scale device, the energy

Figure 28 Nanofabrication challenges compares a chloroplast which is one of natures photonic devices with a charged-coupled device (CCD), which is a man-made photonic device.

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transfer mechanism used within the structure is also unique. The transfer of photonic energy through the structure is carried out by the Forster photonic energy transfer mechanism. The photonic energy transfer process, while not an internal reflection process, can still in some sense be thought of as the equivalent of molecular fiber optics. Thus, such a mechanism could potentially be very useful for communication between molecular and nanoscale components and devices. To date, it has been difficult to design a synthetic model of a solid-state photonic transfer system with any of the properties or efficiency of the biological systems. Some of the problems with enabling this type of nanofabrication is the lack of a suitable ‘‘process’’ to carry out the integration of molecular and nano-scale components into higher order devices and systems. Thus one key issue for nanotechnology is the need for suitable nanofabrication processes and technologies. 10.7.2

Electric Field Array Approach to Micro/Nanofabrication

Active microelectronic DNA arrays have to date been developed primarily for applications in genomic research and DNA diagnostics. These active microelectronic devices have the ability to create a variety of reconfigurable electric field transport geometry on the array surface. This capability allows charged reagent and analyte molecules (DNA, RNA, proteins, enzymes), nanostructures, cells, and even micron-scale structures to be moved to or from any of the microscopic test sites (microlocations) on the device surface. When specific DNA hybridization reactions are carried out on the array, the device is actually using electric fields to direct the self-assembly of DNA molecules at the specified test site or microlocation on the chip surface. Microelectronic arrays have been used to demonstrate the organization of complex fluorescent DNA molecular structures and mechanisms within selected microlocations on the array device (capture probes, target DNA sequences, and reporter probes). Thus, in principle these active devices serve as a semiconductor ‘‘host board’’ array for the nanofabrication of DNA-derivatized component molecules into more complex structures. The DNA molecule, with its intrinsic programmable and selfassembly properties, can be derivatized with a variety of molecular

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electronic or photonic donor or acceptor groups. DNA molecules can also be attached to larger nanostructures, including metallic nanoparticles and organic nanospheres (nanotubes), and to microstructures, including metal and silicon dioxide surfaces. In essence, these active microelectronic host board arrays can allow one to carry out an ‘‘electric field pick-and-place’’ process for the heterogeneous integration of molecular, nano-scale and micron-scale components into more complex two- and three-dimensional structures within the defined perimeters of larger silicon or semiconductor structures (Fig. 29). The use of electric field–assisted self-assembly technology encompasses a broad area of potential applications from nearer-term heterogeneous integration processes for photonic and microelectronic device fabrication, to the development of high-density optical storage materials, to the longer-term nanofabrication of true molecular electronic circuits and devices. The electric field–assisted self-assembly technology is based on three key physical principles: (1) the use of

Figure 29 Scheme for an ‘‘electronic pick-and-place’’ process which uses a microelectronic ‘‘host board’’ array to carry out the heterogeneous integration of molecular, nano-scale, and micron-scale components into higher-order devices and systems.

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functionalized DNA structures as ‘‘Lego’’ blocks for nanofabrication; (2) the use of DNA as a selective ‘‘glue’’ that provides intrinsic selfassembly properties to molecular and nanoscale electronic components and structures (carbon nanotubes, organic molecular electronic switches, etc.); and (3) the use of an active microelectronic array for electric field–assisted self-assembly of any modified electronic/ photonic components and structures into integrated circuits and devices. A potential near-term application of the ‘‘electronic pick-andplace’’ process is for the heterogeneous integration of various microfabricated lift-off components (lasers, diodes, etc.) into more highly integrated photonic/electronic devices (displays, arrays, etc.). Figure 30 shows a specially designed 20-µm light-emitting diode (LED) lift-off device and host board microelectronic array which is designed to carry out the electric field transport of the LED devices to a specific microlocation on the array where the device is attached and metal connections

Figure 30 Diagram of a specially designed 20-µm light-emitting diode (LED) lift-off device, and a specially designed microelectronic host board array which is used to transport and attach the LED to a selected microlocation on its surface.

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Figure 31 Series of images that show the electric field driven transport of a 20-µm LED across the array surface to the selected microlocation where it is subsequently attached and activated.

are made. Figure 31 shows the transport sequence of the LED being moved by electric fields to the specified microlocation, where it will become attached and the proper metal connections made. Figure 32 shows the attached LED being activated and producing light emission. The attachment process involves making the proper in situ metal connections between the p and n regions on the device and the host board. These metal connections are shown in more detail in Figure 33. This figure shows a portion of the tin/lead (Sn-Pb) metal bonding material between the LED and the attachment microlocation on the array surface. A more detailed description on using microelectronic arrays for the transport and attachment of LED structures is discussed by Edman and coworkers [28,29]. Other potential application of the electronic pick-and-place process would be to enable the development of DNA chromophore based high-density optical storage materials. Chromophoric DNA polymers

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Figure 32 Images which show the attached and annealed (bonded) LED, and the activated LED emitting light.

Figure 33 Scanning electron microscope images of the attached and bonded LED, showing a section of the Sn-Pb bonding material connecting the LED to the array surface.

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can be designed to absorb light energy at a single wavelength and reemit at predetermined multiple wavelengths [33]. This DNA chromophoric material has the potential for high bit density optical storage that can be read out as frequency and intensity of the emitted light. In the longer term, the electronic pick-and-place process will be used to carry out the nanofabrication of highly integrated molecular electronic circuits and devices. There are now a relatively large number of research groups involved in the development of molecular electronic components, but only limited ways to carry out the large-scale integration of these molecular and nano-scale components into useful devices. The microelectronic array based electronic pick-and-place process could have key advantages over conventional pick-and-place heterogeneous integration processes. These advantages would include: (1) 4 orders of magnitude size scaling (micron scale ↔ molecular scale); (2) high-fidelity component recognition and self-organization imparted by DNA; and (3) use of an electric field to provide selected transport and orientation of components. Thus, electric field–assisted heterogeneous integration technology has the hierarchical logic. This hierarchical logic (shown in Fig. 34) allows one to control the organization and communication of structures and components from molecular level self-assembly → submicron self-assembly → (Micron Scale 3D Integrated Structures and Devices) ← micron-scale devices ← from macroscopic scale lift-off device fabrication. This concept brings together the best aspects of a pick-and-place process and self-assembly processes to create a potentially viable nanofabrication process. 10.8 CONCLUSIONS Active microelectronic array technology provides a number of distinct advantages for DNA hybridization analysis, as well as for other affinity-based assays that are important for genomic and molecular biology research and for clinical diagnostics. Microelectronic arrays have been designed and fabricated with from 25 to 10,000 microscopic test sites. The higher test site density devices have CMOS elements incorporated into the underlying silicon structure that provide on-board control of current and voltage to each of the test sites on the device. Microelectronic chips are incorporated into a cartridge-type device so as to be

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Figure 34 A micro/nanofabrication process for controlling the organization, assembly, and communication of structures and components from molecular level to the submicron or nano-scale level to the microscopic or micronscale level.

conveniently used with a probe loading station and fluorescent detection system. Active microelectronic arrays are differentiated from other ‘‘passive’’ DNA chip or array technologies by a number of important attributes. Active microelectronic arrays allow DNA molecules (oligonucleotide probes, target DNA/RNA sequences, PCR amplicons, etc.) to be rapidly transported and selectively addressed (spotted) to any of the test sites on the microelectronic array surface. Electronic hybridization allows DNA molecules (probes, targets, amplicons, etc.) to be rapidly transported and selectively hybridized at any test site on the microelectronic array. The large concentration effect achieved by the electric field leads to a significant increase in DNA hybridization rate at the selected test sites. Hybridization which might take hours on a conventional array requires only a minute or two using electronic hybridization. Additionally, electronic hybridization can be carried out under substantially denaturing conditions, which provides many additional

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advantages. The electric field can also be used to provide electronic stringency control, which improves selectivity and allows rapid discrimination of single-base mismatches in point mutation and SNP genotyping analysis. In general, microelectronic arrays have proven extremely useful for carrying out genotyping on difficult or problematic point mutations and single-nucleotide polymorphisms (SNPs). In the case of short tandem repeats (STRs), electronic hybridization is one of the only viable array hybridization methods used for this type of polymorphism. In many of these cases electronic hybridization appears to overcome some of the more intrinsic problems of the basic hybridization process itself. In addition to hybridization analysis, microelectronic arrays have proven useful for carrying out immunoassays, on-chip DNA amplification, and complex cell separation procedures. Microelectronic arrays can be integrated into miniaturized sample-toanswer systems that may ultimately be used for a number of point-ofcare applications. In the longer term, microelectronic arrays may also be used for nanofabrication processes to carry out the heterogeneous integration of molecular, nano-scale, and micro-scale components into useful electronic and photonic devices and systems. REFERENCES 1. Fodor, S.P.A., Read, J.L., Pirrung, M.C., Stryer, L., Lu, A.T., and Solas, D., 1991. Science, 251:767–773. 2. Fodor, S.P., Rava, RP., Huang, X.C., Pease, A.C., Holmes, C.P., and Adams, C.L., 1993. Nature, 364:555–556. 3. Chee, M., Yang, R., Hubbell, E., Berno, A., Huang, X.C., Stern, D., Winkler, J., Lockhart, D.J., Morris, M.S., and Fodor, S.P.A., 1996. Science, 274:610–614. 4. Eggers, M., Hogan, M., Reich, R.K., Lamture, J.B., Ehrlich, D., Hollis, M., Kosicki, B., Powdrill, T., Beattie, K., Smith, S., Varma, R., Gangadharan, R., Mallik, A., Burke, B., and Wallace, D., 1994. Biotechniques, 17:516–524. 5. Lamture, J.B., Beattie, K.L., Burke, B.E., Eggers, M.D., Ehrlich, D.J., Fowler, R., Hollis, M.A., Kosicki, B.B., Reich, R.K., Smith, S.R., Varma, R.S., and Hogan, M.E., 1994. Nucleic Acids Res., 22:2121– 2125.

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Heller, M.J., O’Connell, J.P., and Krihak, M., 2001. Anal. Chem., 73: 1549–1559. Cheng, J., Sheldon, E.L., Wu, L., Heller, M.J., and O’Connell, J.P., 1998. Anal. Chem., 70:2321–2326. Edman, C.F., Swint, R.B., Gurtner, C., Formosa, R.E., Roh, S.D., Lee, K.E., Swanson, P.D., Ackley, D.E., Coleman, J.J., and Heller, M.J., 2000. IEEE Photonics Technol. Lett., 12:1198–2000. Edmen, C.F., Gurtner, C., Formosa, R.E., Coleman, J.J., and Heller, M.J., 2000. High Density Interconnects, 3:30–35. Pethig, R., and Markx, G.H., 1997. Trends Biotechnol., 15:426–432. Markx, G.H., Huang, Y., Zhou, X.F., and Pethig, R., 1994. Microbiology, 140:585–591. Gracias, D.H., Tien, J., Breen, T.L., Hsu, C., and Whitesides, G.M., 2000. Science, 289:170–172. Heller, M.J., and Tullis, R.H., 1991. Nanotechnology, 2:165–171.

11 The Flow-thru Chip A Miniature, Three-Dimensional Biochip Platform Adam Steel, Matt Torres, John Hartwell, Yong-Yi Yu, Nan Ting, Glenn Hoke, and Hongjun Yang Gene Logic, Incorporated, Gaithersburg, Maryland

11.1 INTRODUCTION Integrated, microfabricated device (IMD) manufacturing is a rapidly developing multidisciplinary field. The concept of fabricating miniaturized molecular assay systems, so called labs-on-a-chip, that are capable of performing thousands of analyses simultaneously has its roots in the electronics and semiconductor chip industry. The arrangement of electronic components that make up a circuit board are replaced by an arrangement of microscopic plumbing, valves, thermistors, and molecular recognition elements on an IMD. There is a great deal of interest in IMD technology not only because of the variety of applications such as chemical and biological analysis, diagnostics, and forensics, but also because of the variety of sciences and technologies that are involved in the development process. IMD development routinely draws upon such diverse fields as material science, physical chemistry, biochemistry, nucleic acid chemistry, molecular biology, electrical and mechanical engineering, optics, image and data analysis, database management, and laboratory automation. Development of convenient technologies for quantitative determination of nucleic acids and immunoproteins is 271

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of high value to the scientific, medical, diagnostic, biotechnology, and pharmaceutical communities. As IMD research continues to expand it is very likely that more diverse fields will be tapped for contributions and will incorporate and benefit from the technology. A theme in IMD development is to combine and control as many labor- and reagent-intensive processes as possible inside a single device that is inexpensive and easy to use. In most cases, IMD production is not simply a matter of reducing the physical dimensions of existing measurement systems, interfacing the various components is a particular challenge. A fully functional IMD contains several components, each with its own function. Complete systems for a particular analysis are being developed. For example, a microchip device that incorporates cell lysis, PCR amplification, and electrophoretic sizing has been reported in the literature [1]. Alternatively, development of individual IMD components are being developed by niche groups. As examples, novel structures for sorting and detection of macromolecules which can be integrated into a device have been developed by independent groups [2,3]. At Gene Logic, we are developing an improved sensor platform in which molecular interactions occur within the three-dimensional volumes of ordered microchannels rather than at a two-dimensional surface. The technology has been termed the Flow-thru Chip [4]. While development of this technology has been focused on a standalone readout mechanism for nucleic acid expression analysis to date, integration of the technology as a component of an IMD is a clear possibility. Arrays for biomolecule analysis have evolved from the dot-blot on a nylon membrane to the microarray on a microscope slide. Currently available microarray technologies still require a significant extent of manual manipulation to complete an assay. With the continuing evolution and miniaturization of heterogeneous assay systems, a number of system integration possibilities are apparent. With respect to the integration of a DNA chip assay platform, we have identified three major areas for potential improvement: fluid transfer, chip transfer, and process timing. Several fluid transfer steps (e.g., prewashing, blocking, sample injection, hybridization, flushing, staining, and postwashing) are required to complete an entire microarray assay. Automation of the fluid transfer steps has not been readily performed in macroscopic

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systems. Chip transfer steps are needed between shakers, fluid-handling systems, incubators, and detection devices. Precise timing of each step is difficult to achieve in nonintegrated systems. The benefit of implementing automated control to the entire process such that active user intervention is not necessary was designed into the Gene Logic Flowthru Chip system. The Flow-thru concept is illustrated in Figure 1. Microchannels connect the upper and lower faces of a chip in such a manner that fluid can flow through the chip. Probes, nucleic acids, or antibodies, for example, are deposited into one or more discrete microchannels of the chip to create a microarray that is used to carry out multiple determinations in parallel. The term probe will be used to describe a species which is immobilized on the surface of the microchannels and has some specific interaction with a ‘‘target’’ that is part of a fluid test mixture. The microarray application is just one of many application in

Figure 1 Conceptual schematic of the Flow-thru Chip. The chip is composed of an ordered array of microscopic channels that transverse the thickness of the substrate. Arrays of probes are deposited on the chip in spots that incorporate several microchannels. Fluid flows through the chip and biological recognition reactions occur within the confined volumes of the microchannels.

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which an advanced biochip could be integrated as a component of an IMD. The Flow-thru Chip can also be used in a variety of ways such as a microreactor, concentrator, and microcuvette, as examples. The contents of this chapter provide an introduction to the Flow-thru concept, a technical overview of chip manufacture and implementation, and a survey of a few notable applications in development.

11.2 THE FLOW-THRU CHIP SYSTEM 11.2.1

The Flow-thru Concept

Consider the mechanics of a heterogeneous bimolecular reaction, that is, a reaction involving two species, one immobilized at a surface and the second present in a fluid in contact with that surface. Probes and targets can be, as examples, small molecules or drugs, nucleic acids, proteins, or even cells. In conventional hybridization chambers the bulk of the target-containing volume is a considerable distance, on the molecular length scale, from the reaction site on the surface. For this case, mass transport of target to the immobilized probe represents the ratelimiting step in all but the slowest molecular interactions. The diffusion coefficient for DNA molecules is small, typically ⬍1 ⫻ 10 ⫺7 cm 2 sec and decreases with target length, so that the capture rate is slow unless significant convective currents are introduced to enhance mass transport [5]. Microarray fabrication involves spatial distribution of individual probes. Hence, for the number of determinations per chip to increase either the spot size must decrease or the dimensions of the chip must increase. In the former case, mass transport issues are exacerbated by increasing the chip size. In the latter, smaller spot size translates to a smaller number of probes which in turn limits the breadth of the dynamic range for the measurement. Sensitivity to targets at spots of decreasing dimensions may become limited also due to the dependence of DNA binding on the concentration of the immobilized probe [6–8]. An approach to enhance chip performance is to increase the surface area by using a three-dimensional support matrix. The third dimension has been implemented on chips in the form of a gel pad and, as we shall discuss at length in this chapter, the Flow-thru Chip [9,10].

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Chip Materials and Properties

Attributes desired in Flow-thru Chip solid supports include ease of manufacture, ease of processing, mechanical strength, and cost. The chip material should allow efficient immobilization of probes, either directly or through an intermediate surface coating, and should have sufficient tensile strength and chemical resistance to withstand binding reaction conditions. In practice, Flow-thru Chip materials contain a high density of noninterconnected channels that connect the upper and lower faces of the chip. The diameter and packing density of the microchannels can be varied to match the requirements of specific applications. The microchannels provide increased surface area. The increased surface area is a function of the microchannel radius, the open area ratio, and the chip thickness. For channels with 10 µm diameter, packed at a 50% open area ratio, the capillary array provides roughly 100 times the surface area of a flat chip. The surface area enhancement increases for thicker chips with smaller, more densely packed microchannels. However, the physical dimensions that provide the largest surface area may not be workable in practice. In particular, the pressure required to flow liquid through the chip increases dramatically as the microchannel diameter is reduced [11]. Also, reliable quantitation of signals from chips of increasing thickness becomes quite difficult due to the optical properties of the microchannel matrix [12]. Two chip materials have been used in the production of Flowthru chips to date: glass capillary arrays and electrochemically etched porous silicon. Development using the glass chips is more advanced at this stage and the bulk of data discussed in this chapter was generated using glass capillary arrays. Work with the porous silicon material is in the early stages of characterization. Other materials that have been examined for the Flow-thru Chip include porous metal oxides and template-assisted porous polymers [13]. Glass Capillary Arrays Microporous glass is commercially produced for a variety of applications that are quite different from biochips (e.g., as intensifiers in night vision goggles). The material is known as microchannel plate glass or glass capillary arrays. Glass capillary arrays consist of many parallel

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tubes fused together in a uniform and mechanically rigid matrix. Capillary arrays are commercially available with microchannels ranging from 2 µm to ⬎50 µm, and array thickness from 0.1 mm to several millimeters. An image of a glass capillary array with 10-µm channels at an open area ratio of 50% under 100 times magnification is given in Figure 2. A 5-nL spot of fluorophore solution is seen in the field. The porous glass has reasonable tensile strength and chemical resistance. Microarray development on glass is facilitated by the large body of research that has been devoted to immobilization of biomolecules onto glass, and robust attachment strategies have already been developed [14,15]. While quite convenient for biochip development in a research environment, the glass capillary arrays are not as suitable for direct inclusion in an integrated device. The material is not readily machined, and producing glass microchannel structures within a larger ensemble would be quite difficult. The optical properties of the capillary array glass provide an additional limit to the materials usefulness. Crosstalk between channels, seen as the diffuse halo around the spot in Figure

Figure 2 Phase contrast and fluorescence images of a 5-nL spot of dye solution at 100⫻ magnification. The channel diameters are 10 µm on both materials. The contrast has been adjusted in each of the images.

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2, makes spots appear broader than the actual dimensions. The channels act as waveguides, funneling photons to the edges of the chip. The waveguiding property of the material is observed by noting that the interior of the channels is significantly brighter than the interstitial regions. In spite of the noted limitations, glass capillary arrays have been used with notable success. Macroporous Silicon Silicon has many potential advantages as a support matrix for biochips and IMD fabrication. Silicon’s compatibility with a host of analytical techniques for surface analysis has resulted in tremendous knowledge of materials and methods to control its surface properties. Silicon presents a pristine, well-ordered surface in comparison to glass, which is amorphous and quite heterogeneous. The formation of microchannel arrays with high aspect ratios by electrochemical etching was reported in 1990 [16]. The channels are arrayed in patterns predetermined by an initial photolithography step. However, there are a limited number of patterns that are accessible to the electrochemical etching process. The combination of pattern and microchannel diameter produces a constant porosity material during efficient etching [17]. Diameters in the range of 0.3–20 µm can be produced with aspect ratios of up to 250, meaning that the process easily produces 10-µm channels across a 500µm-thick silicon wafer. An image of a 5 nL spot of fluorescent solution on a porous silicon chip with 10-µm channels is included in Figure 2. The fluorescence image presented in Figure 2 for the silicon chip was acquired with a longer integration time than used for the glass chip. The emission from the silicon chip is roughly an order of magnitude less than the glass chip for the equivalent loading of fluorophore. Quenching of emissions from within silicon microchannels presents a particular challenge to biochip development on this material. Silicon is a very workable material via micromachining and lithographic methods and has already been used in IMD production [18]. Microchannel-containing silicon regions can be integrated into traditional silicon constructs quite easily using photolithographic methods [13]. Methods to produce stratified structures in bulk silicon are being developed that will greatly increase the possibilities for device integra-

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tion of a flow-through component. In addition, robust chemistries for modification of the silicon surface have been introduced recently that will permit facile immobilization of biomolecules [19,20]. 11.2.3

Chip Preparation

The primary objective in chip preparation is to permanently immobilize probes in a functional conformation such that the appropriate target can bind to the probe and nonspecific binding of other components in the target mixture is reduced to a level that provides useful signal-tobackground ratios. Biomolecule immobilization can be accomplished by adsorption, entrapment, and covalent attachment among others [15]. Covalent attachment is the preferred method for ‘‘permanent’’ immobilization; however, great care must be exercised when developing a covalent bonding scheme to ensure that the activity of the probe is not adversely effected. Whereas few organic functional groups react directly with an activated silica or crystalline silicon surface, an intermediate layer is commonly used to enhance biomolecule immobilization. Functionalized organosilanes have been used extensively as an intermediate layer on glass and silicon chips [14,15]. Biomolecules are immobilized to the intermediate layer using crosslinking chemistry. Several extensive reviews of bioconjugate techniques are available; see for example the text by Greg Hermanson of Pierce Chemical Company [21]. Methods for Flow-thru Chip Preparation Effective cleaning of the support matrix prior to silanization is extremely important to reproducible chip production. The cleaning process should both remove contaminates (e.g., oils, dirt, and detergents) and, in the case where silanization is to be used, generate reactive hydroxyl groups on the silica surface [15]. Chips were cut to 12-mm 2 dimensions from bulk glass and silicon wafers using a diamond scribe. Glass was cleaned by sequential sonication in 1 N nitric acid, absolute ethanol, and deionized water for 10 min each. Following the water rinse, glass chips were baked at 80°C for at least 4 hr to remove the liquid from the capillary array. Silicon chips were cleaned using warm piranha (70% concentrated sulfuric acid, 30% hydrogen peroxide;

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WARNING—piranha reacts violently with organic solvents), followed by sonication in deionized water for 10 min twice. A thermal oxide was grown on the porous silicon by heating to 1300°C in wet oxygen. Solution phase silanization was used for all chip preparations. Chips were immersed in a 2% solution of mercaptopropyltrimethoxysilane (MPTS; Aldrich Chemical Company, St. Louis, MO) in dry toluene (Fisher Scientific, Pittsburgh, PA) for 1 hr. Excess silanization solution was removed using blotting paper, followed by 10-min serial sonications in dry toluene, a toluene–absolute ethanol mixture (50/50 by volume), and absolute ethanol. Silanized chips were baked at 80°C for 16 hr to ensure complete hydrolysis of the methoxy residues. The number of mercaptan groups introduced to glass chips was determined using a thiol quantitation kit from Molecular Probes (Eugene, OR). The available mercaptan surface coverage is 2.6 ⫻ 10 13 groups/cm 2 prior to the baking step. Baking the chips results in a decrease in available mercaptan surface coverage to 2.0 ⫻ 10 13 groups/cm 2, presumably due to oxidation of the mercaptan to the sulfinate or sulfonate. Chips typically are spotted within hours of removal from the baking oven and stored in a dehumidified, light-tight container until used. Nucleic acid probes were synthesized (Research Genetics, Huntsville, AL) with a primary amine at the 3′ end of the sequence (Glen Research, Sterling, VA). Proteins were immobilized through free primary amines in the lysine residues in the amino acid sequence. Probes were reacted with a crosslinker in 1⫻ saline sodium citrate buffer (Sigma Chemical Company, St. Louis, MO). The reaction was allowed to proceed for 1 hr at room temperature prior to deposition on the chip. Aliquots, 5 nL, of the reaction mixture were spotted onto MPTS silanized chips using a Packard BioChip piezoelectric spotter (Packard Instrument Co., Meriden, CT). Nucleic acid probes were deposited at a density of 1.4 ⫻ 10 13 probes/cm 2, less than the number of potential mercaptan binding sites, 2.0 ⫻ 10 13 SH/cm 2 from above. Fluorescentand isotope-labeled probes were used to determine the binding efficiency of amine-modified nucleic acid segments on the MPTS surface. Probes were typically retained at the surface in the 30–60% range. The immobilized probe density in each spot was on the order of 6 ⫻ 10 12 probes/cm 2. Literature values for probe surface densities on silanized glass surfaces range from 10 12 to 10 13 probes/cm 2 [6,9,22]. This range

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of probe surface densities has been reported optimal for DNA hybridization [8,23,24]. Spotted chips were allowed to dry in a dehumidified, light-tight box for 24 hr prior to blocking. Spotted chips have been stored for as long as 6 months prior to use without observing significant loss of performance. In protein binding assays, monoclonal mouse antihuman IgG subclass antibodies (Fc specific), polyclonal goat antihuman IgG (H ⫹ L specific) antibody (Zymed, San Francisco, CA), monoclonal mouse antihaptoglobin, anti–bovine serum albumin (Sigma, St. Louis, MO) and antihuman interleukin-2 (PharMigen, San Diego, CA) were immobilized via a crosslinker in 1⫻ SSC buffer at room temperature. Aliquots, 25 nL, of the antibody/crosslinker mixture were deposited on a MPTS silanized Flow-thru Chip using a sapphiretipped needle on a Hamilton Microlab 2200 at a 0.8-mm pitch. Immediately after Flow-thru spotting, the chip was transferred to a highhumidity chamber for 40 min to complete immobilization of the capture antibodies to the chip. Antibody chips were stored at 4°C until used. Nonspecific adsorption of target can create a high level of background signal which in turn reduces the sensitivity of the assay. Different blocking protocols have been followed for nucleic acid and protein experiments. For nucleic acids, chips were incubated in 250 µL of a blocking solution for 15 min and baked at 80°C for 1 hr. Blocked chips were stored in a dehumidified, light-tight container until used. For proteins, chips were blocked by incubation in blocking buffer containing 10% fetal bovine serum for 1 hr at room temperature. Blocked chips were blotted dry and stored at 4°C for several days prior to use. Flow-thru Chip Hybridization and Detection Prepared chips were mounted in a specially designed cartridge described below prior to running an assay. The cartridge mounts into a fluid-handling system that contains injection loops for the target mixture and a staining dye where appropriate. Prior to injection of the target mixture, buffer was run through the chip and flow cell for 15 min to remove any loosely bound material in the microchannels from probe deposition and blocking. Nucleic acid hybridizations were conducted in 5 ⫻ SSPE buffer. Antibody reactions were performed in phosphate buffered saline (100 mM NaCl, 100 mM sodium phosphate). The flow

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rate was 0.2 mL/min and reactions were conducted at room temperature unless otherwise stated. Fluorescence intensity measurements were determined using a Nikon Eclipse E800 (Tokyo, Japan) microscope fitted with a Nikon PCM-2000 confocal accessory and a Hamamatsu C4742-95 Orca monochrome camera (Hamamatsu City, Japan) for image capture. The Orca camera has a 1280 ⫻ 1024 pixel CCD chip with 6.7 ⫻ 6.7 µm pixels, three binning modes, and a 12-bit A/D converter. Image capture and analysis were performed using SimplePCI, a software application that provides image capture, enhancement, and analysis in an integrated package (Compix, Cranberry Township, PA). 11.2.4

Advantages of Flow-thru Chip Geometry

Development of the Flow-thru Chip to this point has focused on demonstrating that the three-dimensional structure of microchannel containing materials can be exploited to provide enhanced biochip performance. The potential advantages of developing biosensors in these materials include: (1) improved responsiveness and dynamic range due to the increased surface area compared to a flat surface geometry; (2) reduced assay times due to enhanced mass transport within the channels; and (3) more uniform probe deposition and higher array densities due to wetting properties of microporous materials. A series of experiments which show that the advertised advantages can be realized in a Flow-thru Chip are summarized in the following sections prior to discussion of the Gene Logic integrated Flow-thru Chip system. Increased Responsiveness and Dynamic Range Assay responsiveness is given by the slope of the observed signal versus analyte concentration curve and dynamic range by the analytical signal range over which the response curve is linear. The amount and distribution of binding sites in a spot are determining factors for assay responsiveness and dynamic range. The larger the number of probes within an analysis area, the greater the responsiveness to target, more signal per-unit concentration, and the higher the binding capacity for target. The number of probes that can be immobilized in a given analy-

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sis area is a function of the chip surface area. The additional surface area gained by adding depth to the chip rather than expanding the lateral dimensions in the Flow-thru Chip is the basis for increasing the responsiveness and dynamic range. The enhanced performance of a Flow-thru Chip compared to a flat chip is evidenced by the results in Figure 3. In these experiments chips were prepared by deposition of equivalent volumes of equimolar probe stocks on a microchannel glass chip and a flat glass slide. Both of the support matrixes were treated with MPTS silane. Probe attachment was very similar on the chips as determined by the level of immobilization of a fluorescently tagged probe. Hybridizations were carried out in a Gene Logic Flow-thru Chip cartridge and were monitored in situ. The target was 50 nM and was recirculated around the chips at 0.2 mL/min using the scheme detailed below. The increase in fluorescence,

Figure 3 Comparison of hybridization signal on the Flow-thru Chip and a traditional flat substrate. The average fluorescence signal to background ratio (n ⫽ 3) is plotted versus the hybridization time. The fluorescence was measured in situ during recirculation of 50-nM target. Filled circles represent Flow-thru Chip signal, empty circles flat glass signal, and lines are provided as a guide to eye only. The signal intensity is roughly 40 times brighter and the hybridization rate is roughly six times faster on the Flow-thru Chip than on the flat glass.

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corrected for background, from a single spot is given as a function of the hybridization time in the figure. The plateau values on the two chips are substantially different. The signal reached on the flow-thru chip is ⬎40 times that of the flat glass chip. As stated above, the enhancement in responsiveness and dynamic range on the Flow-thru Chip is a consequence of the larger surface area for probe immobilization. Based on the ratio of the surface areas for the Flow-thru and flat chips, a 100fold enhancement was expected in this experiment. The observed increase in dynamic range is smaller than the theoretical enhancement presumably due to optical limitations associated with the three-dimensional nature of the Flow-thru support matrix [12]. Enhanced Reaction Rates Reactions on microarrays typically require significant incubation periods not because the reaction kinetics are slow but because it takes a long time for targets to find the complementary probe on the array. Transport of slow-moving target molecules in the large reaction volume, relative to the molecular length scale, is the rate-limiting step for the majority of biomolecular reactions in macroscopic systems [23,25,26]. To enhance target transport to flat chips convective fluid currents are often introduced by shaking, turning, etc., or by dramatically shrinking the dimensions of the assay. However, at a boundary layer of fluid remains unperturbed at the surface on the order of 100 µm thick independent of any mechanical efforts to effect mixing [27]. Physically confining the probe and target to as small a volume as possible enhances the rate of reaction [28,29]. In the current flow-through geometry, the target solution flows through microchannels with a diameter that is significantly smaller than a typical boundary layer thickness at planar chips. The microscopic diameter of each microchannel results in a large surface area-to-volume ratio. Hence, binding events that occur within the small volume of the microchannels do so with high efficiency. Bringing the reactants close together results in efficient transport due to diffusion. The residence time of target molecules in the microchannels is controlled by the bulk fluid flow rate through the chip. The bulk flow rate can be adjusted such that the residence time in the microchannels is long relative to the time needed for target to migrate the short distance to the channel wall via diffusional transport. The

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enhanced mass transport of target to reactive site increases the rate of hybridization by speeding up the rate-limiting step. The relative rate of reaction on a Flow-thru Chip and a flat chip is indicated by the data in Figure 3. The slope of the signal-time curves for each chip geometry provides a measure of the transport of target to the probe spots on each type of chip. The slope for the Flow-thru Chip is six times that of the flat glass. Diffusional mass transport scales with the square of distance. Taking the boundary layer thickness at a planar chip as 100 µm and a microchannel radius of 5 µm, the expected ratio of the slopes should be 400 [27]. However, the preceding calculation fails to account for lateral migration of targets which is allowed in the flat case but not in the Flow-thru Chip. A correction factor based on the reaction volumes for each geometry is needed to make a reasonable prediction of the relative rates of reaction on the two geometries. The ratio of the slopes in Figure 3 should correspond to the cube root of the diffusional mass transport calculated ratio when the dimensionality of the geometries is considered. Using this method the calculated ratio is 7.5 for the assumed lengths which agrees quite well with the observed value. Uniform Probe Distribution and Increased Spot Density The microchannels that make up a Flow-thru Chip behave like capillaries; thus, liquid is drawn into the volume of the chip by capillary action. The wetting behavior provides the practical advantage of more homogeneous probe distributions than on flat chips. The extent to which the capillaries control the wetting of chips is visible in Figure 2, where the packing geometry of the microchannels determines spot shape. On the glass capillary array, with the microchannels packed in a hexagonal lattice, the spot appears hexagonal. On the silicon chip, with the microchannels packed in a square lattice, the spot appears octagonal. In each case the spot shape is the best approximation of a circle that is possible on the given lattice structure. Two distinct advantages of the Flow-thru Chip result from the capillary wetting behavior: (1) probe immobilization is facilitated due to slower evaporation of small (nanoliter) droplets of probe deposition solution; and (2) higher-density arrays are possible because the same volume of liquid has a smaller footprint on the Flow-

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thru Chip than on a flat chip. With regard to the former advantage, rapid evaporation of probe deposition solution may result in inconsistent immobilization of the probes on the surface. Loss of solvent may not provide sufficient time for proper reaction of immobilization chemistries with the surface producing nonuniform probe immobilization. The distribution of probes within the spot is important because the ability of target to bind to probe is a function of the probe surface density [6,8]. When probes are assembled too densely, the binding reaction is inhibited due to steric interactions. Also, nonuniform distribution of the probes in the spot can give rise to irregular intensities, which complicates image analysis. Spot densities on flat glass chips can reach densities of roughly 800 spots/cm 2. In the chip configurations used to date, equivalent volumes deposited on a microchannel containing chip have a 60% smaller diameter than on a flat chip. The smaller spot size permits higher array densities, as much as three times higher, on a Flow-thru Chip. 11.3 FLOW-THRU CHIP INTEGRATED SYSTEM Consider the several operations required to complete an entire microarray assay. There are fluid transfers, chip transfers, and incubations, all of which need to be performed as reproducibly as possible. Fluid transfers in a common assay include prewashing, blocking, sample injection, hybridization, flushing, staining, and postwashing. Each step requires a user to apply fresh solutions to the array because the actions are not readily automated. Chip transfer between shakers, fluid handling systems, incubators, and detection devices are also common. Precise timing of each of the operations is difficult to achieve in nonintegrated systems but is highly desirable to ensure reproducible results. The benefit of implementing automated control to the entire process such that active user intervention is not necessary was designed into the Gene Logic Flow-thru Chip system. In the current system configuration the sample volumes and fluid handling systems are macroscopic. However, future implementations of the technology may employ flow cells and fluid handling mechanisms engineered to use smaller sample volumes using a integrated device approach.

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Fluid delivery to the chip is a primary concern for implementation of the Flow-thru Chip geometry. There are a number of factors that have been considered in the design of the fluid interface to the chip. First, the diameter of the channels in the Flow-thru Chip support matrix is generally small, on the order of 10 µm, such that fluid flows through the chip only when a pressure gradient is present across the chip. The diameter is so small that fluid flow through the microchannels is laminar, Reynolds number ⬍⬍2300. Because a pressure gradient is necessary to get fluid flow through the chip, sealing on the edges of the chip becomes rather important. Improper sealing results in a fluid shortcircuit around the edge of the chip and little or no flow through the chip. Second, reproducible chip performance requires uniform fluid flow across the entire chip face. Nonuniform flow results in nonhomogeneous array performance due to differences in local fluid distribution. Third, a means to redistribute the target solution is desirable for detection of low abundance targets. Once a target enters a channel, it cannot laterally migrate into other channels until it has come out of the interior of the chip and the probability that a target enters a channel containing a complementary probe is small. There are roughly 1 million microchannels in a 1-cm 2 chip and a single probe spot contains around 100 microchannels. Hence, recirculation of the target mixture through the chip increases the probability that a particular target has the opportunity to bind with the complementary probe. Fourth, the fluid interface should permit visual access to the chip for observation by the detection device. The Flow-thru Chip fluid delivery system consists of two major components: a cartridge that guides the movement of fluid through the chip, and a fluidics station that addresses fluid delivery to and from the cartridge. The cartridge is the heart of the system. The system has been designed such that a chip, once loaded, is not manipulated directly by a user at any point in the assay. Also, the cartridge design includes a window to the chip which permits real-time imaging, a feature not commonly associated with microarrays. The fluidics station is the processing unit of the fluid delivery system, designed to direct the necessary buffers, target solutions, and staining solutions through the cartridge. The fluidics station design has been heavily driven by the concept of sample recirculation to boost assay sensitivity.

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The Flow-thru Chip Cartridge

The cartridge is machined from anodized aluminum with two separate fluid chambers: a reaction chamber containing the Flow-thru Chip, and a thermal fluid chamber. Process or sample fluid enters the cell from underneath the chip, flows up through the chip, and then out through an exit above the chip. A compression fit window is located on the top of the cartridge above the chip to permit viewing in situ. Viton gaskets are used to seal the chip in the cartridge. Experiments to determine the uniformity of fluid distribution are described below. The thermal fluid chamber houses a heated water stream just beneath the reaction chamber to control the temperature in the reaction chamber. Each chamber is connected to the fluidics station via a pin and septum.

Single-Pass Assays Flow-thru Chip assays are fast because both reactants are confined to a small volume inside the microchannels. Under appropriate flow rates, targets are captured from solution by the complementary probe during each pass through a microchannel on the chip. Because not all of the microchannels on the chip contain a probe to capture targets, the sensitivity of the assay depends on the total number of microchannels in the chip and the number of microchannels that contain probe. The sensitivity of nucleic acid hybridizations in the Flow-thru Chip for a single pass of target mixture through the chip has been tested. The signal intensity is plotted versus the target concentration in Figure 4. The total active chip area is 1 cm 2. Each chip contained three identical probe spots. The probe was an 18mer, designed to be complementary to a fluorescently labeled 65mer target beginning at the 5′ end. For each concentration represented in the figure, 15 pM to 1 µM, 85 µL of target solution was passed through the chip at a flow rate of 0.2 mL/min. This flow rate provides a residence time in the channels of 8 sec. The signal for the 1-nM target concentration is two standard deviations above the background level. For concentrations ⬎1 nM the signal increases linearly with a slope very close to unity, representing three orders of magnitude of dynamic range. For nucleic acid analysis nanomolar-level sensitivity is sufficient for only a few applications. Hence,

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Figure 4 Response of the FTC to single-pass hybridization of target. Volumes of 85 µL at target concentrations from 15 pM to 1 µM were passed through a chip a single time at a flow rate of 0.2 mL/min. Data points represent the average of three determinations per chip.

some form of amplification, either target or signal, is necessary for most applications. The sensitivity of the single-pass assay is in some ways limited by the geometry that has been presented. In the 1-cm 2 active area chip each spot represents roughly 0.0001 of the total number of microchannels. Therefore, assuming uniform fluid distribution, the fraction of target that passes through a channel containing a complementary probe is quite small. When the partitioning of the target mixture is taken into account, the single-pass assay sensitivity of 1 nM detected roughly 10 attomoles of target bound to the probe spot. Shrinking the dimensions of the active chip surface is one means of boosting the sensitivity of the single-pass assay; another is to recirculate the target through the chip as will be discussed below. In a recirculation scheme, the assay sensitivity should scale with the number of passes through the chip. Few examples of microarrays developed for parallel analysis of proteins have been reported even though the use of microarrays for nucleic acid analysis is of great interest [13]. The Flow-thru Chip has been used in single-pass mode for the determination of clinically rele-

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vant levels of human immunoglobulins, the four IgG subclasses in particular, via a sandwich immunoassay. Normal serum levels in human blood range from 0.5 to 9.0 mg/mL for each of the IgG subclasses. Chips were prepared for assays as described above. In the single-pass Flow-thru Chip assay, 100 µL of 10 µg/mL total human IgG (IgG1, 65% by mass; IgG2, 25%, IgG3, 6%; IgG4, 4%) in PBS buffer was passed through the chip at 0.2 mL/min. Nonspecifically and loosely bound proteins were removed by washing with PBS buffer (pH 7.0) at 0.4 mL/min for 5 min prior to injecting the target mixture. The chip was stained for readout by passing detection antibody (100 µL of 100 µg/mL FITC-labeled monoclonal mouse antihuman IgG [Fab specific]; Sigma, St. Louis, MO) in PBS buffer through the chip, followed by a final wash with PBS buffer. An image of the chip response is given in Figure 5. Capture antibodies for each of the IgG subclasses is present as are three negative controls (haptoglobin, bovine serum albumin, and interleukin-2) and a positive control (polyclonal IgG). The relative intensities on the Flow-thru Chip, 4 ⬎ 3 ⬎ 1 ⬎ 2, are quite similar to the relative intensities observed in a colorimetric microtiter plate assay (4 ⬎ 3 ⬎ 2 ⬎ 1). The relative intensities are determined by the affinity between the capture antibody and the target IgG, which is different for

Figure 5 Fluorescence antibody detection of IgG subclasses on the Flowthru Chip. An 85-µL aliquot of 10 µg/mL total human IgG was passed through a chip a single time at 0.2 mL/min. The antibodies were measured using a fluorescent-labeled monoclonal mouse antihuman IgG (Fab-specific) antibody.

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each subclasses. The signal-to-background ratio for IgG 4 is close to 100 for the data in the figure. Little nonspecific binding was observed on negative controls. Designing an assay to determine the presence or absence of immunoglobulins provides limited information. In many cases it is more valuable to measure diagnostically relevant changes in IgG subclass abundance. Greater than fourfold changes in subclass abundance have been reported as diagnostically relevant for certain diseases in the literature [30]. For example, low IgG 2 and/or low IgG 3 levels were found in patients with juvenile diabetes. The ability of the Flow-thru Chip to discriminate fourfold changes in subclass abundance was determined by mixing purified human IgGs. The response when the abundance of a single subclass was reduced fourfold while other subclass levels were unchanged was determined for IgG 2 and IgG 3. In both cases, the chip response was reduced to 20% of the ‘‘normal’’ value, indicating that the Flow-thru Chip is sensitive to at least fourfold changes in a single IgG subclass abundance compared to a normal distribution. 11.3.2

The Flow-thru Chip Fluidics Station

As discussed above, single pass of the target through the chip has limited sensitivity, requiring some form of amplification to determine lowabundance targets. Sensitivity can be increased by passing the target mixture through the chip several times. A prototype fluidics station has been built to implement target recirculation through the chip cartridge. In its simplest form, the fluidics station is composed of three valves, a peristaltic pump, and a Flow-thru Chip cartridge as shown in the schematic in Figure 6. The peristaltic pump drives fluid throughout the system that is connected in a fluid path. Peristaltic pumping is noninvasive, can pump fluid in a closed loop, and is easily configured to a large range of fluid flow rates. Alternative pumping schemes such as syringe pumps or diaphragm pumps are not compatible with a recirculating fluid delivery system. Of the three valves, one is used for buffer selection, the second to isolate a recirculation loop which includes the peristaltic pump and the cartridge, and the third for sample injection. The recirculation valve is open (i.e., set to waste) during pre- and postwashing steps of a hybridization and closed during the reaction step, creating a closed loop, the target recirculation loop, which includes the chip.

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Figure 6 Schematic representation of the Gene Logic fluidics station. The configuration depicted here contains three valves (buffer selection, recirculation control, and sample injection), a peristaltic pump, and the chip cartridge. The recirculation valve switches between creating an open circuit from the buffer reservoirs to the waste and a closed circuit including the pump, the samples injection valve, and the chip cartridge.

The sample injection valve is only open (i.e., set to inject) during the reaction and during a staining step where applicable. Integrated Bread Board Unit The integrated bread board unit (IBBU) is a prototype instrument that integrates fluid transfer steps, chip transfer steps, and process timing for Flow-thru Chip hybridization assays. A picture of one IBBU unit is shown in Figure 7. Each unit has five independently programmable fluid modules each of which accommodates a single cartridge. A retractable lid is attached to each module, one of which is up in the figure, to limit photobleaching of fluorescent markers and provide thermal stability during the hybridization process. Each fluid module is designed to operate in a manner similar to the simple scheme outlined above; however, an additional injection loop was incorporated into the IBBU as a dedicated staining port. The entire system is controlled by a personal computer running Lab-View software (National Instruments, Austin, TX). Flow rates, temperature, buffer selection, and timing for each fluid module are controlled independently. A status window updates the progress of each module and includes start, pause, and stop

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Figure 7 Picture of the Gene Logic Flow-thru Chip integrated bread-board unit (IBBU). Each unit contains five independently programmable fluid modules for Flow-thru Chip hybridization.

toggles. The toggles permit manipulation or removal of the cartridge during an experiment in the event of an technical problem. A total of four valves (Valco Instruments, Houston, TX) are included in the system for buffer selection, sample injection, recirculation, and staining. The pump head (Watson-Marlow Bredel, Wilmington, MA) is capable of flow rates from 600 to 1400 µL/min using marprene tubing (tubing supplier, Watson-Marlow Bredel, Wilmington, MA). A single set of buffer reservoirs services all of the modules. Two temperature baths are incorporated to provide temperature control during each of the process steps. Thermal fluid is distributed to each of the modules from a common line using an external pump (Tuthill, Concord, CA) where it passes through the thermal fluid chamber of the cartridge and through a preheating tube that surrounds the process fluid tubing prior to the cartridge. The reaction temperature can be controlled between ambient and 50°C at a resolution of 1°C. Uniform Fluid Distribution Reproducible assay performance is highly dependent on the uniformity of fluid distribution across the chip face. To test the uniformity of fluid flow, a microarray pattern was spotted covering 60% of the viewable chip area. The array consisted of 16 identical 4 ⫻ 4 subarrays. Each subarray contains four different probes, denoted as probe 1 through 4, spotted in quadruplicate. An image of a uniformity test array run in a Gene Logic Flow-thru cartridge is given in Figure 8. The target mixture

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Figure 8 Uniformity of hybridization to microarray on a chip in the Flowthru Chip cartridge. The microarray contains 256 spots, divided into 16 identical 4 ⫻ 4 arrays, covering 0.6 cm 2 of the chip. In each 4 ⫻ 4 array the columns contain the same probe sequence. A target mixture containing 2 nM corresponding to probe 1, 10 nM to probe 2, 20 nM to probe 3, and no target to probe 4 (from left to right) was hybridized to the array for 4 hr in recirculation mode at 200 µL/min. The response is nearly linear with concentration, and the average standard deviation for each probe type response is roughly 10%.

used to test the array contained targets complementary to probes 1, 2 and 3. The concentrations for targets 1, 2, and 3 were 2 nM, 10 nM, and 20 nM, respectively. The target to probe 4 was left out as a control. The recirculation flow rate was 0.2 mL/min for a total hybridization period of 4 hr, corresponding to roughly 50 cycles through the chip recirculation loop. The standard deviations are 14%, 12%, and 7% for probe/target pairs 1, 2, and 3, respectively. The small variability for each probe/target pair indicates that the fluid distribution is quite uniform over the entire microarray. The standard deviations are noticeably smaller if the spots at the edge of the array are removed from the statistical sample set or by imaging subsections of the array individually.

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These observations suggest that the reported standard deviations are inflated as a result of the detection system rather than an indicator of nonuniform fluid distribution. In alternate flow cell geometries ‘‘hot’’ and ‘‘cold’’ spots develop on the chip along prevalent fluid flow lines. The standard deviations for a single probe on the uniformity test array can be as high as 50% in such alternate flow cells. The relative signal intensities on the chip are 1, 3.2, and 8.5 for probe/target pairs 1, 2, and 3, respectively. The analytical response on the Flow-thru Chip is nearly linear with various target concentrations; expected relative intensities were 1, 5, and 10 from the solution concentrations. The signal intensity does scale linearly with target concentration when a single probe/target pair is tested. Hence, we postulate that the relative intensities on the uniformity chip were a reflection of the difference in probe/ target pair affinities. In Situ Imaging A unique property of the Flow-thru Chip fluid delivery system is the ability to view reactions in situ. DNA hybridizations were monitored for fluorescent-labeled targets in real time by mounting the chip cartridge assembly on an epifluorescence microscope. As sample passes through the chip, specific targets are captured from solution by the probes on the chip. Under the recirculation condition, target accumulates over time resulting in greater fluorescence intensity. Similar experiments with flat glass arrays are possible. As discussed above, realtime monitoring was used to show that hybridization on flat glass chips occurs at a much slower rate (⬃6⫻ slower) and the final signal-tonoise ratio of individual spots is lower by roughly 40 times than on a Flow-thru Chip [12]. Real-time hybridization detection can be quite useful for assay optimization, DNA melting studies, and kinetics-based gene expression analysis. The in situ observation capability may also integrate well with Lab-on-a-Chip sample amplification platforms [1,31] whereby rapid analysis of a relatively pure PCR amplicons is desired. Temperature Control of Hybridization There is significant interest in using DNA chips to determine single nucleotide polymorphisms (SNPs) for diagnostics [18,31]. SNPs are

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single-base mutations that can contribute to diseases. Mapping of SNPs is currently being performed to determine the role in disease development and progression. Once determined, the SNP can be used as a diagnostic marker for testing individuals [32]. Within the possible combinations of nucleic acid sequence homology, successful discrimination between perfectly matched sequences (PM) and a single base pair mismatch (SBMM) is the most difficult to detect. Discrimination between the PM and SBMM requires control of the conditions, or stringency, during hybridization. Stringency parameters, temperature, and ionic strength are commonly determined empirically for a given PM and SBMM pair. The ability to discriminate PM and SBMM sequences was investigated on the Flow-thru Chip system by varying the temperature during hybridization for a series of PM and mismatch probes. Flow-thru Chips were prepared with three different 18mer probes including a PM, SBMM, and a three-base mismatch (TBMM), following the procedure outlined above. The probes were complementary to a 65mer sequence based on a segment of beta-actin mRNA (GenBank M17851). The target was was modified with a Texas Red fluorescent reporter group to permit direct detection on the chip. The probes align to the first 18 bases of the target from the 5′ end. The SBMM was created by inserting a pyrimidine for a purine, creating a pyrimidinepyrimidine mismatch, at the 10th nucleotide on the PM sequence. The TBMM was created by similar modification at nucleotides 9–11 on the PM sequence. Chips were hybridized to 1.2 moles of labeled target for 2 hr at 700 µL/min flow rate. Using the IBBU, fluid temperature inside the cartridge and recirculation loop is controlled during prehybridization washing and hybridization. Hybridizations were performed at 20, 25, 28, 35, 40, and 46°C (⫾2°C). Figure 9 contains a plot of the fluorescence intensity for the PM, SBMM, and TBMM probe sets versus the hybridization temperature. The signal for the TBMM is initially below the other probes and falls off dramatically as the hybridization temperature is increased. The SBMM and PM show comparable results up to 35°C at which point the SBMM signal starts to decline at a faster rate than the PM signal. The chip results are in qualitative agreement with solution melts for the probe/target pairs. The T m’s for the PM, SBMM, and TBMM in solution are 73, 63, and 49°C, respectively. The chip data indicate that the hybridization temperature on the chip

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Figure 9 Chip signal as a function of hybridization temperature for a series of probes with no (PM, solid symbol, solid line), one (SBMM, open symbol, solid line), and three (TBMM, open symbol, dash line) mismatched nucleotides in the sequence. The error bars represent 1 SD on three determinations per temperature.

need not reach the solution T m temperature to provide the same level of discrimination. The data also indicate that near complete complementarity of probe and target is necessary to result in hybridization on the chip. Control of the temperature within well-defined limits during hybridization was engineered into the IBBU system and provides notable benefit for nucleic acid analyses, including SNPs. Analysis of Gene Expression Patterns A potential application of the Flow-thru Chip is drug screening. The viability of the chip as a readout mechanism in a screening assay was investigated using the MCF-7 cell line. MCF-7 cells are estrogenresponsive breast adenocarcinoma cells that have been used previously as a model system for breast cancer drugs [33]. Initial studies have examined the effect of an antiestrogen, tamoxifen, on the estradiol response of MCF-7 cells. Tamoxifen treatment of cells inhibits the induction of several genes that occur upon estradiol treatment. It is thought

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that tamoxifen inhibits proliferation of estrogen-responsive cells through competitive binding to the estrogen receptor. The Flow-thru Chip has been used as an analysis platform for a subset of genes that have been reported in the literature as differentially regulated in the MCF-7 system under estradiol and tamoxifen treatment. The experimental protocols for estradiol and tamoxifen treatment have been described in detail elsewhere [33]. Briefly, MCF-7 (ATTC, Manassas, VA) cells were maintained in MEM alpha medium supplemented with 10% heat-inactivated fetal bovine serum, penicillin-streptomycin, and 10 µg/mL bovine insulin (complete medium) (Life Technologies, Gaithersburg, MD). For the treatment experiment 1 ⫻ 10 5 cells/cm 2 were plated into T75 flasks, using 2 mL/10 cm 2 complete medium and allowed to attach for 24 hr. During the next 3 days the cells were rinsed twice with PBS and incubated with MEM phenolfree medium supplemented with 5% dextran sulfate/charcoal stripped fetal bovine serum, penicillin-streptomycin, L-glutamine, 10µg/mL bovine insulin, and 1⫻ MEM nonessential amino acids (stripped medium). The stripped medium was changed daily for 2 more days. Four samples were prepared: a nontreated control, an estradiol treatment, a tamoxifen treatment, and a mixed estradiol and tamoxifen treatment. The cells were then incubated for 24 hr with stripped medium in the presence of 10 nM 17-beta-estradiol and a combination of 10 nM 17beta-estradiol with 1 µM tamoxifen. The total RNA was harvested by removing the media and applying the Trizol reagent kit according to manufacturer’s instructions (Life Technologies, Gaithersburg, MD). The purity and integrity of the total RNA was assessed by the 260/ 280 and the 28s/18s ratios. The total RNA was converted into cRNA. As before, biotins were incorporated into the cRNA via NTPs for detection and targets were fragmented to create nucleic acid segments in the range of 25–50 nucleotides. Flow-thru Chips were prepared with probes for 12 genes, including four controls. Two 18mer probes were spotted per gene and each probe was spotted in triplicate. Chips were hybridized to 8 µg of fragmented cRNA for 12 hr at 200 µL/min flow rate. Hybrids were stained with a streptavidin–Texas Red conjugate for 15 min and imaged using the Nikon microscope. Quantitative RTPCR measurements were also made to verify the changes in expression levels for each of the monitored genes across the treatments.

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The results for selected genes are presented in Figure 10. Absolute correlation of signal intensities from two separate images is not generally performed; rather, signal intensities between images are compared via normalization through a reference or control. In Figure 10, the signal intensity for each of the two probes per gene is given as the ratio of the intensity for the indicated treatment sample versus the intensity for the nontreated control sample. Hence, genes that are upregulated produce a signal ⬎1 and downregulated genes a signal ⬍1. The data for one signal normalization control gene and two test genes are represented in Figure 10. The control gene is glyceraldehyde 3-phosphate dehydrogenase (GAPDH, G) and the test genes are Cathepsin D (C), BRCA-1 (B). GAPDH was assumed to be present at equivalent levels in all samples because it has been reported to be unaffected by the

Figure 10 Change in signal intensity for selected genes in estradiol and estradiol ⫹ tamoxifen treated MCF-7 cells relative to untreated control. The genes included are GAPDH (G), Cathepsin D (C), and BRCA-1 (B). The designations of 1 and 2 correspond to individual probe sequences for the targets. The error bars represent the standard deviation from six independent determinations per probe.

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indicated treatments in this study [33]. Quantitative RT-PCR data indicate that GAPDH was not differentially expressed between the samples. The RT-PCR data indicate sixfold induction for Cathepsin D and fourfold induction for BRCA-1 in the estradiol-treated samples. The tamoxifen- and estradiol-treated samples displayed only twofold induction of Cathepsin D and no change in BRCA-1 expression relative to the control sample. The chip data from Figure 10 showed the same trend as the quantitative RT-PCR for these genes; the signal for each gene was increased in the estradiol treatment and inhibited for the estradiol and tamoxifen treatment. However, the signal response was compressed on the chip compared to the RT-PCR measurements. For example, instead of a sixfold increase for Cathepsin D, the signal increased by only 60% over the control level. Compression of signal intensities on microarrays has been documented previously in the literature [34,35]. Possible causes for the compression include saturating the probe spot with target, taking the signal ratio with one of the measurements near the lower limit of detection, and nonlinear hybridization of different probe/target sequence pairs. Continued investigation of the viability of the chip as a readout mechanism in a screening assay is being conducted with a larger gene set, drug set, and temporal dosage regiment. 11.4 CONCLUSION The Flow-thru Chip was introduced as an advanced analysis platform, with distinct advantages over traditional chips for biological sensing applications. Materials and methods of Flow-thru Chip analysis were presented, and the advantages gained by using an organized threedimensional material for biomolecular analysis were demonstrated for microarray applications. In particular, nucleic acids were determined at subnanomolar concentrations within a few minutes and immunoglobulins were analyzed at diagnostically relevant levels with a single pass through the Flow-thru Chip. Integration of the Flow-thru Chip to a fluid delivery system has provided significant improvements for this biochip platform. An expressly designed cartridge and fluid-handling station was engineered to minimize user interaction during the hybridization assay resulting in more reproducible results with increased sam-

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ple throughput. While currently a macroscopic system, miniaturization, and integration of flow-through components into more advanced systems are envisioned. ACKNOWLEDGMENTS We would like to thank our collaborators for their invaluable input. Professor Anthony Cass, Dr. Catherine Halliwell, and Dr. Christophe Valet of Imperial College (London) for issues regarding silanization of glass and silicon; Professor Jonathon Cooper and Vincent Benoit of Glasgow University (Glasgow) for optical studies of flow-through chips and developing methods to produce microchannels in silicon; Larry Millstein for the concept diagram in Figure 1 and critical reading of the manuscript; and Dr. Kenneth Beattie and Dr. Mitch Doktycz of Oak Ridge National Labs for discussion of the Flow-thru Chip. REFERENCES 1. Waters, L.C., et al. (1998) Microchip device for cell lysis, multiplex PCR amplification, and electrophoretic sizing. Anal. Chem. 70:158–162. 2. Duke, T.A.J. and R.H. Austin. (1998) Microfabricated sieve for the continuous sorting of macromolecules. Phys. Rev. Lett. 80:1552–1555. 3. Delemarche, E., et al. (1998) Microfluidic networks for chemical patterning of substrates: design and application to bioassays. J. Am. Chem. Soc. 120:500–508. 4. Beattie, K.L. (1996) Microfabricated, flowthrough porous apparatus for discrete detection of binding reactions. U.S. Patent 5843767. 5. Tinland, B., A. Pluen, J. Sturm, and G. Weill. (1997) Persistence length of single-stranded DNA. Macromolecules 30:5763–5765. 6. Bamdad, C. (1998) The use of variable density self-assembled monolayers to probe the structure of target molecules. Biophys. J. 75:1989–1996. 7. Beier, M., and J. Hoheisel. (1999) Versatile derivatisation of solid support media for covalent bonding on DNA-microchips. Nucleic Acids Res. 27:1970–1977. 8. Steel, A.B., T.M. Herne, and M.J. Tarlov. (1998) Electrochemical quantitation of DNA immobilized on gold. Anal. Chem. 70:4670–4677. 9. Doktycz, M.J., and K.L. Beattie. (1997) Genosensors and model hybrid-

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12 DNA Arrayed on Plastic Devices Robert S. Matson Beckman Coulter, Inc., Fullerton, California

12.1 INTRODUCTION The advent of DNA array technology as a tool for genetic analysis represents one of the new paradigms for processing biologically significant data in a massively parallel fashion. The original intent of the technology was for solid-phase genetic sequencing (so-called SBH or sequencing by hybridization), but this has given rise to both tools for gene discovery and diagnostic applications. The advantages of the ‘‘reverse’’ blot (genosensors, type II array format) over the conventional dot blot (Southern blot, type I format) or other competing technologies such as the polymerase chain reaction can be argued. However, the most fundamentally distinct advantage has been that of platform minaturization which in turn has led to a reduced sample size as well as the ability to perform higher throughput analysis using automation tools. It has also been the advances in solidphase substrate chemistries that have led to the successful implementation of array-based technologies. The earliest contributors to the notion and utility of array based hybridization include the work of Saiki et al. [1] on HLA genotyping and Dattagupta et al. [2] on micro-organism detection. These groups relied upon nylon membranes as the solid support. Southern et al. [3] were the first to demonstrate the in situ synthesis of oligonucleotide 303

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probes using glass plates as the solid phase; Pease et al. [4] provided an elegant demonstration of photolithographic-based in situ synthesis of oligonucleotides on silicon chips. Less complex arrays were created for diagnostic applications such as described by Matson et al. [5,6] and Wenhert et al. [7] and Weiler and Hoheisel [8] on plastic substrates. Khrapko et al. [9] attached probes within a polyacrylamide gel matrix deposited on glass. Sosnowski et al. [10] immobilized probes within the permeation layer that coated electronically addressable electrodes in a microfabricated chip device, and Schena et al. [11], in order to analyze for gene expression, very simply created microarrays of cDNA by printing onto polylysine-coated glass microscope slides. Thus, a number of approaches are available for the construction of arrays and there are a variety of solid-phase materials from which to choose. From a manufacturing viewpoint the selection of a solidphase support requires that the materials and processes be inexpensive, physically durable, and chemically compatible, and utilize simple chemistries while maintaining good ligand specificity. In this chapter we will discuss only the utility of array technology based upon plastic as the solid support. 12.2 PLASTICS AS SUBSTRATES There are today a very large number of different polymer and polymer blends that we commonly refer to as plastics that are available for use in the manufacture of household, medical, scientific, and industrial products. Common plastics used in the biomedical field include polystyrene, polycarbonate, and polypropylene. These thermoplastics can be processed in roll form, melt-blown, or injection molded into useful shapes such as test tubes and microtiter plates. Polypropylene (PP) is among the most chemically inert and physically durable plastics. The plastic is not easily fractured, has high tensile strength, and is quite resistant to a variety of solvents, strong acids, and bases. It is also compatible with strong oxidizers and chaotropic agents. These physical and chemical attributes are of great advantage, especially for work involving harsh chemical environments such as employed during DNA synthesis where strong organic solvents and oxidizers are required. Polypropylene materials are also suitable as

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hybridization supports having low nonspecific adsorption of biologicals such as proteins and nucleic acids, and with good optical properties such as low intrinsic fluorescence. Few solid supports can satisfy all of these criteria to be useful for DNA synthesis, hybridization, and detection. For these reasons polypropylene was selected for in situ synthesis of oligonucleotide arrays. We will also describe its use in the form of molded array carriers for use on an automated robotic workstation. 12.3 SURFACE MODIFICATION To be useful as a DNA synthesis support, surface reactive groups must be introduced from which to tether the growing oligonucleotide chain.

Figure 1 Free radical plasma mechanism.

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Owing to polypropylene’s inertness it was necessary to create these surface reactive groups using a chemical gas plasma process. A chemical plasma is a highly reactive gas containing free radicals, ions, electrons, and other metastable species. Plasmas are created during the discharge of electromagnetic radiation—e.g., sun spots, lightning bolts. In our case, a radiofrequency (RF) plasma generator set at 13.56 MHz was used to create a plasma discharge (RFPD; radiofrequency plasma discharge) in an evacuated chamber containing ammonia gas. The vacuum process permits work at or near room temperature. Free radicals within the plasma collide with the plastic substrate creating surface free radicals. The plasma reaction generally takes place within the first 10– ˚ of the surface. The mechanism is described in Figure 1. This 1000 A process converts approximately 2–5% of the surface to amine functionalities and as such does not appreciably affect the bulk properties of the plastic. The aminated surface is uniform and stable. The amine density for films processed from a continuous roll RFPD is 5–6 pmoles/mm2, corresponding to an oligonucleotide probe array density of 1–3 pmoles/mm2. Loading can be varied during the plasma amination process by adjusting gas partial pressure and RF power.

12.4 ATTACHMENT OF DNA 12.4.1

In Situ Oligonucleotide Array Synthesis System

Once aminated the films can be stored for several years if properly sealed and protected from ultraviolet light sources. Standard β-cyanoethyl N,N-diisopropyl (CED)-phosphoramidite DNA synthesis conditions can be employed to create ‘‘nucleotide polypropylene’’ [5]. The first cycle of this reaction is described in Figure 2. The result is the creation of oligonucleotides tethered to the surface 3′ → 5′ by a very stable phosphoramide bond. However, to create in situ arrays a special DNA synthesizer is required having multiport switching valves that direct synthesis chemicals into a microchannel reactor block. In our case, an early prototype of the Oligo 1000 DNA Synthesizer (Beckman Coulter, Inc.) was converted into a 64-line dispenser that delivered syn-

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Figure 2 The first cycle of the nucleotide polypropylene reaction.

thesis reagents to a 64-channel block reactor. A central eight-port valve was used to distribute chemicals to four (16-port) valves that interfaced with the block (Fig. 3). The activated polypropylene film was sandwiched between two aluminum plates. One plate was machined with 64 500-µm-diameter grooves, 0.8 mm depth, 62 mm length, with a 0.8 mm centerline separation between channels. The polypropylene film served as the sealing gasket with the activated side facing the grooved channels and the untreated side facing the top plate. When properly clamped the reactor block and the plastic sheet provided a leak-free environment with no crosstalk between channels. Then synthesis reagents were pumped through the channels permitting reaction of these reagents with only the activated side of the sheet. The entire process was controlled under LabView software using a Macintosh computer.

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Figure 3 An 8-port valve distributes chemicals to interface with the block.

The CED-phosphoramidite base monomers (A, C, G, T) were delivered in serial to the channel block according to the directions given by a run file. For example, the A monomer phosphoramidite was delivered only into those channels where the next base to be coupled to the growing polymer (probe) were A. Once all of the A sites were addressed, the C channels would be filled, followed by G and T couplings. An 8 cm ⫻ 8 cm sheet synthesis of 64 different oligonucleotide sequences in 500-µm-wide strips across the film could be accomplished in an overnight run. Probes up to 98mer have been synthesized and demonstrated to be of good quality. More typically, however, probe sequences ranged from 9mer to 18mer for routine work. Verification of coupling efficiency was accomplished by monitoring the release of the trityl cation from the support. Efficiencies were typically ⫹98%. The quality of the oligonucleotide could also be determined by introducing a cleaveable link (e.g., succinate spacer) to the support prior to synthesis.

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Figure 4 Checking for purity with capillary electrophoresis.

The cleaved oligonucleotide could then be checked for purity using capillary electrophoresis (Fig. 4). Upon completion of synthesis the film sheet is removed from the block and bulk processed for base deprotection in ammonia. Following a water rinse the film is ready for hybridization experiments. In most instances, the oligonucleotide array was sectioned into ‘‘dipsticks.’’ Each of these was approximately 0.5 cm ⫻ 7.5 cm and contained the 64 probe sequences in rows (Fig. 3). Lane-to-lane variation obtained from hybridization analysis is estimated to be at a 7–10% coefficient of variation (CV). It was also possible to interrupt synthesis and remove the plastic sheet, rotate it 90°, and continue with a second round of synthesis. This creates 500-µm-square features (Fig. 3) in a 64 ⫻ 64 array or 4096 probe sequences that are arranged into 64 families. Thus, an array containing 64 different 6mers created by the first synthesis round would serve as leader sequences for a set of 64 second-round sequences of 9mers to create 15mers arranged in 64 families according to the leader sequence. For example, consider two channels (Ch 1 and Ch 2) used to synthesize leader sequences 3′GTACGT5′ and 3′AAAAAA5′, respectively. In the second round Ch 1 now calls for the synthesis of 3′GGGCCCTTT5′ and Ch 2 ⫽ AGGCCCTTT, giving rise to the following probes:

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Matson Resulting probes

Ch 1, 1st round GTACGT ⫹ Ch 1, 2nd round GGGCCCTTT⫽ GTACGT GGGCCCTTT Ch 1, 1st round GTACGT ⫹ Ch 2, 2nd round AGGCCCTTT⫽ GTACGT AGGCCCTTT Ch 2, 1st round AAAAAA ⫹ Ch 1, 2nd round GGGCCCTTT⫽ AAAAAA GGGCCCTTT Ch 2, 1st round AAAAAA ⫹ Ch 2, 2nd round AGGCCCTTT⫽ AAAAAA AGGCCCTTT

This is repeated for all 64 channels. In this manner, a 4096-element cystic fibrosis array based upon a single allele was created to demonstrate the process and serve as a model for predicting hybridization efficiency [6]. In other studies, Wehnert et al. [7] created an oligonucleotide array strip of 60 trinucleotide (21mers) and four dinucleotide (20mers) tandem repeats to screen for the presence of STRs (short tandem repeats) in PCR fragments and cosmids. Using a modified synthesis chemistry, Weiler and Hoheisel [8] created arrays of cleavable probes on polypropylene film for use as sequencing primers. Although the design of the in situ synthesis system permitted only limited flexibility in the number and placement of probes within an array, the ability to create a series of probe sequences with such a rapid turnaround time has some advantage in determining an optimal probe design. For example, one can vary a single probe sequence and it’s mismatch over 64 permutations of 3′ to 5′ position with length and select the optimal probe from the set from hybridization analysis the next day. Thus, we were able to design a series of probes for ras point mutations within a narrow Tm range (e.g. 54.3 ⫾ 2.6°C) that avoided unwanted loop structures that could be formed between complementary regions of neighboring probes (interstrand duplexes). 12.4.2

Quality Control

A criticism certainly of in situ synthesized arrays has been in the assessment of the quality of probes synthesized at each location on the substrate surface. Ideally, one would also like to measure the number of probe molecules tethered. Aside from radiolabeling, methodology that is nondestructive for the array and that can be used on a routine basis to quantitatively determine single probe density is currently unavailable. The qualitative characterization of arrays is possible. In our in situ system, the routine inspection of the quality of the synthesis was

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assessed by staining individual strips or the full sheet. Owing to the probe density of in situ synthesized arrays, common DNA stains could be employed. A nondestructive stain, SYBR Green I (Molecular Probes, Inc.), was used to determine the completeness of synthesis in channels (probe rows). Documentation could be provided by using a CCD camera system and a UV lightbox. Following staining the strips could be cleared of the stain in methanol, rinsed, and used directly for hybridization. However, because SYBR Green I gives faint signals, it has to be viewed at 254 nm. This could potentially lead to the crosslinking of adjacent probes to each other if viewed over an extended time (e.g., 5–10 mins). There was also some sequence-dependent variation; e.g., rows containing sequences rich in dA were more faintly stained than others. For these reasons, statistical sampling of strips was more often employed for quality control purposes using an InVitrogen stain. This stain (DNA Dipstick) uniformly stains all rows with a visible blue dye. However, in this case, the staining is destructive and prevents use of these strips for hybridization. 12.4.3

Postimmobilization Processes

As previously mentioned, the in situ synthesis system described here is most useful in probe design and for exploratory experimentation where only a limited number of probes are required. To create more complex arrays that are cost-effective it is advisable to use a postattachment process. Thus, once optimal probes are selected these are then synthesized in bulk on a conventional (cleavable) DNA synthesis support and postattached to activated plastic to create arrays. For example, probes can be spotted down onto plastic substrates using either a noncontact process such as piezoelectric dispensing or by contact printing using solid or split-pins with microarrayers. To provide for efficient coupling to plastics, a new activation chemistry has been developed based upon the use of diethylamine sulfur trifluoride (DAST) reagent. As shown in Figure 5 the plastic is derivatized with surface carboxyl groups that react with DAST to form the highly reactive acyl fluoride. 5′ amino oligonucleotide probes are then spotted down on the substrate and become covalently linked to the surface in a matter of seconds. This is followed by a capping

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Matson

Figure 5 Plastic is derivatized with surface carboxyl groups that react with DAST to form the highly reactive acyl fluoride.

(quenching) of residual groups by soaking in ethanol for ⬃2 hr. The array is then rinsed with water ready for use in hybridization experiments. In addition to oligonucleotides, cDNA can be immobilized by this process. Based upon staining, the oligonucleotide probe density is less than that which can be achieved by in situ synthesis. For postattachment, oligonucleotide probes are prepared at 20 µM and picoliter volumes are delivered to the surface. Assuming near quantitative coupling efficiency, a probe density based upon 200-µm-diameter features would be at ⬍1 pmole/mm2 as compared to the in situ synthesized arrays which have been determined to have densities in the range of 1–3 pmoles/mm2. 12.5 TARGET LABELING STRATEGIES One of the disadvantages of using arrays has been the issue of probetarget hybridization efficiency. Owing to steric hindrance caused by probe surface crowding, it is difficult to achieve levels of hybridization much above 1% of the probe density using larger DNA targets (e.g., 200–5000 bp). One approach has been to use shorter fragments of ssDNA or RNA as targets. For example, hybridization of labeled cRNA fragments. These are created by in vitro reverse transcription of DNA into labeled and subsequently fragmented cRNA. This is the strategy employed in using the Affymetix’s GeneChip [12].

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Another approach and one that we have adopted is that of signal amplification. In this case, target DNA is either primer labeled, e.g., 5′ biotin primer, or internally labeled, e.g., biotin-dCTP incorporation during amplification. Hybridized target is detected using a streptavidin conjugate such as with alkaline phosphatase or horseradish peroxidase. We have used the alkaline phosphatase-ELF substrate (enzyme-labeled fluoresence detection; Molecular Probes, Inc.) signal detection with internally labeled DNA targets to achieve a level of sensitivity at a solution concentration within the low pM to fM range. For example, the overnight hybridization of G3PDH cDNA at 60°C resulted in detection at 2 fM ⬃ 200,000 copies of cDNA in 200 µL hybridization buffer. It should also be noted that not all signal generation systems work well with plastic substrates. Since the surface remains largely uncharged, the wetting of the surface and adsorption of materials based upon ionic interaction mechanisms are more difficult. Certain chemiluminescent systems that require the sequestering of a light emitting intermediate through charge interaction with the surface do not work well. Signal tends to diffuse from the surface causing a fuzzy image sometimes covering multiple elements of the array. While low nonspecific adsorption of proteins is advantageous for reduced backgrounds, it is difficult to use these as effective blocking agents. In some instances blocking has actually contributed heavily to signal backgrounds. 12.6 AUTOMATION AND SYSTEM INTEGRATION Using a robotic workstation platform (Biomek 2000, Beckman Coulter, Inc.) an automated array hybridization, signal development, and detection system was designed. The system consisted of a standard Biomek 2000 Workstation, a temperature-controlled shaker water bath and a cooled CCD camera system (Fig. 6). The key to the system was an injected molded carrier, called the Biotip, which held an oligonucleotide array (Fig. 7). Ethylenemethacrylic acid copolymer (EMA) with pendant carboxyl groups was injection molded into the form of the Biotip. This material was readily derivatized to the reactive acyl fluoride form for immobilization of the probe. The bottom of the molded piece (⬃ 1 cm ⫻ 1 cm) served as the site for probe attachment. Generally, 10 ⫻ 10 probe arrays of 300-µm features were created for muta-

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Figure 6 The detection system.

Figure 7 The key to the system was an injected molded carrier, called the ‘‘Biotip.’’

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tion screening—e.g., ras oncogene analysis. Printing was accomplished using a BioDot 3000A/D dispenser (Cartesian, Inc.). A 10 ⫻ 10 array can be dispensed to the surface at ⬃1 dot/sec or 1 array in about 50 sec. In addition to oligonucleotide probes, an amino-biotin compound (5′biotinamido pentlyamine, BAPA, Pierce Chemical) was printed as registration markers for use with image analysis software. These BAPA markers also served as a positive control for the enzyme conjugate (streptavidin-alkaline phosphatase) activity. The Biotip could be picked up by a Biomek pipet tool and delivered to different areas of the workstation for automated sample processing beginning with chemical denaturation of the target DNA, neutralization and hybridization, rinsing, addition of signal development reagents, followed by the development and reading of the array by the CCD (Fig. 8). In terms of the Biomek work flow, the process takes approximately 8 min to mix reagents and move Biotips between stations. For a 1-hr hybridization the total time from denaturation through signal read is about 4 hr for 16 samples. One critical issue that developed was the instability of the DNA arrays on the Biotip. The arrays appeared to be stable for only a few weeks. Further investigation determined that substantial amounts of HF (a byproduct of the hydrolysis reaction) were being released out of the plastic. This is believed to be the source of degradation of the DNA on the surface. As a result, acyl fluoride activated EMA has been re-

Figure 8 Biotip array carrier processing.

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placed by polypropylene, which contains low percentage of surface activated acyl fluoride groups. In doing so the DNA array shelf life has been extended to over 1 year. For high-throughput applications the Biotip approach has been replaced by the use of a shallow-well microplate (currently under development) in which probes are printed into the wells of the acyl fluoride activated polypropylene. The BioDot dispenser head is replaced by a split pin print head (Telechem, Inc.) for contact printing. This permits the printing of arrays with 100–400 elements per well. The wells are ⬃ 6.5 mm diameter, 2 mm depth, with a working volume of 25 µL. The microwell array (array of arrays) concept provides for an estimated processing of up to 10 plates (9600 samples) in approximately 15 hr based upon a 3-hr hybridization using a Biomek 2000 equipped with a Gripper tool to move the plates on and off of the workstation. 12.7 SUMMARY As a result of the introduction of new surface modification chemistries, low-to-medium-density DNA arrays can be easily created on molded plastic devices. Such array carriers can be used with currently available robotic workstations. Thus, the use of plastic substrates for array-based technology offers a distinct advantage in scaling up for high throughput mutation screening, gene expression, and diagnostic applications. ACKNOWLEDGMENTS I wish to recognize my colleagues for their work on the development of the array technology described. They are Jang Rampal, Peter Coassin, Raymond Milton, John Fassett, Tom Stachelek, Dave Helphrey, Jack McNeal, and James Osborne (Beckman Coulter, Inc.); Professor Edwin Southern (Oxford); and members of the NIST/ATP-sponsored Genosensor Consortium (especially MIT/Lincoln Laboratories who built the automated CCD camera system). REFERENCES 1. Saiki RK, Walsh PS, Levenson CH, Erlich HA (1989). Proc. Natl. Acad. Sci. USA 86:6230–6234.

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2. Dattagupta N, Rae PMM, Huguenel ED, Carlson E, Lyga A, Shapiro JA, Albarella JP (1989). Anal. Biochem. 177:85–89. 3. Southern EM, Maskos U, Elder JK (1992). Genomics 13:1008–1017. 4. Pease AC, Solas D, Sullivan EJ, Cronin MT, Holmes CP, Fodor SPA (1994). Proc. Natl. Acad. Sci. USA 91:5022–5026. 5. Matson RS, Rampal JB, Coassin PJ (1994). Anal. Biochem. 217:306– 310. 6. Matson RS, Rampal J, Pentoney SL, Anderson PD, Coassin P (1995). Anal. Biochem. 224:110–116. 7. Wehnert MS, Matson RS, Rampal JB, Coassin PJ, Caskey CT (1994). Nucleic Acids Res. 22:1701–1704. 8. Weiler J, Hoheisel JD (1996). Anal. Biochem. 243:218–227. 9. Khrapko KR, Lysov YP, Khorlin AA, Ivanov IB, Yershov GM, Vasilenko SK, Florentiev VL, Mirzabekov AD (1991). J. DNA Sequencing Mapping 1:375–388. 10. Sosnowski RG, Tu E, Butler WF, O’Connell JP, Heller MJ (1997). Proc. Natl. Acad. Sci. USA 94:1119–1123. 11. Schena M, Shalon D, Davis RW, Brown PO (1995). Science 270:467– 470. 12. Wodicka L, Dong H, Mittmann M, Ho H-M, Lockhart DJ (1997). Nat. Biotechnol. 15:1359–1367.

13 Microfluidic Devices Fabricated by Polymer Hot Embossing Holger Becker Mildendo GmbH, Jena, Germany

13.1 INTRODUCTION The commercialization of microsystem technology increasingly requires low-cost microfabrication methods suitable for high-volume production. In many application fields, notably in the life sciences [1], the development of microcomponents or complete microsystems has reached a state where no longer simple demonstrators in various forms of system integration are state of the art, but where complex technology supply chains for volume fabrication have to be built up to address growing markets. The application of miniaturization in fields such as human genome research, of the drug discovery process in the pharmaceutical industry, clinical diagnostics, or analytical chemistry is increasing the performance of conventional methods, and also changes the approach to analytical data generation and handling with concepts like the ‘‘Labon-a-Chip’’ which sits on a doctor’s desk or at the patient’s bedside. In addition, it opens novel markets and generates new business opportunities [2]. The underlying concept of this development is called µ-TAS, the ‘‘total microanalysis system’’ which arose historically from analytical chemistry. Already in the 1970s, in a remarkable effort, Stephen Terry miniaturized a gas chromatography system and integrated the 319

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complete system on a silicon wafer. Tragically, this work [3] went unnoticed for more than a decade, until in 1990 the conceptual µ-TAS paper was published [4]. This article triggered an avalanche of discoveries and developments of planar microfluidic devices on glass or quartz substrates [5–13], which led to a truly exponential growth of this field (for recent reviews, see e.g. [14,15]), first in academic research alone, but since the mid-90s also on a commercial basis. But for an application of miniaturized devices in tasks like highthroughput screening or DNA sequencing, cost is a major issue, which has to be taken into account in the fabrication technology of these devices. Microfabrication technologies that allow the low-cost production of disposable devices on a biocompatible substrate are therefore required. In terms of microfabrication, an ideal biochemical device should have the following properties: 1. 2. 3.

4.

Suitable microfabrication technologies for a large variety of geometries (rectangular, rounded, high aspect ratios, etc.). Low cost for high-volume fabrication (low substrate cost, short fabrication cycle times). Surfaces compatible to the media used without additional treatment (e.g., in microfluidics no sticking or molecular decomposition). For microfluidic applications low conductivity for electrokinetic pumping or electrophoretic separation, low autofluorescence in case of fluorescence detection, and optical transparency.

Polymers as substrate material offer a possible solution to these fabrication challenges and lend themselves to the mass fabrication of microfluidic devices. Their wide range of material properties, their normally low costs, and the development of suitable polymer microfabrication methods like hot embossing over the last years have attracted an enormous interest particularly as this opens up the road to a highvolume production of disposable microfluidic devices, which allows the successful commercialization of the µ-TAS concept. This paper is organized as follows: After a brief introduction into polymer characteristics, the various methods for the microfabrication of the embossing master are examined. After the description of the hot

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embossing process, examples of microfluidic structures are presented. A section on additional process steps (like dicing and bonding) is added to complete the overview on polymer device fabrication. 13.2 POLYMER MATERIALS Polymers are becoming an increasingly important category of materials in microfabrication. Particularly for the life sciences, polymers offer a wide range of suitable material properties as well as the potential of low-cost high-volume fabrication. Polymer replication methods such as hot embossing, injection molding, and casting can be utilized, leading to fabrication costs which allow for disposable devices. Polymers differ in their mechanical properties, optical characteristics, temperature stability, and resistance against chemicals such as acids, alkalines, or organic solutions, and there are also biodegradable polymers, so that there is a polymer material suited for nearly every application. These material aspects are still largely unexploited in microfabrication. 13.2.1

General Properties

Polymers are macromolecular substances with a relative molecular weight between 10,000 and 100,000 daltons and ⬎1000 monomeric units. The polymerization process is started by an initiator substance or a change in the physical parameters (light, pressure, temperature). Due to the great length of the polymer chain, polymers are bulk materials. In most cases polymers are amorphous or in some cases microcrystalline, where the length of the polymer chains is larger than the size of the crystallites. As the chain length of the polymer molecules in bulk material varies, a polymeric material doesn’t have an exactly defined melting temperature. Instead there exists a melt interval, where the viscosity changes strongly and the material turns into a highly viscous mass. The decomposition temperature is another characteristic point above which the thermical cracking of the material starts and the material ceases functioning. Many of these highly molecular substances solidify after cooling under the so-called glass transition temperature T g and the solid phase that results is glass like hard and brittle. For fabrication processes, this is one of the important parameters. If the temperature is increased

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above T g, the material gets plastic-viscous and can be molded. It is important for the molding process to cool the material below T g before demolding. Otherwise, the geometric stability of the molded component can suffer due to the relaxation during the demolding and the resulting entropy elasticity. Softeners can be used to reach lower glass transition temperatures for the respective material. Due to the softeners the elasticity, the impact strength and the expansion of the polymer increase and the hardness decreases. Polymers can be classified into the following three categories according to their molding behavior which is determined by the interconnection of the monomer units in the polymer chain: Thermoplastic Polymers They consist out of unlinked or weakly linked chain molecules. At a temperature above the glass transition temperature, these materials become plastic and can be molded into specific shapes, which they will retain after cooling below T g. Elastomeric Polymers Also very weakly crosslinked polymer chains. If an external force is applied, the molecular chains can be stretched, but relax back to their original state (higher entropy), once the external force is removed. Elastomers also do not melt before reaching their decomposition temperature. Duroplastic Polymers In these materials, the polymer chains are more strongly crosslinked, so a molecular movement for a change in shape is not easily possible. These materials therefore have to be cast into their final shape. They are harder and more brittle than thermoplastic materials and soften only very little before the temperature reaches their decomposition temperature. Meanwhile, a wide variety of polymer materials have been used for microfabrication processes: standard polymer materials such as polyamide (PA), polybutylenterephthalate (PBT), polycarbonate (PC),

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polyethylene (PE), polymethylmethacrylate (PMMA), polyoxymethylene (POM), polypropylene (PP), polyphenylenether (PPE), polystyrene (PS) and polysulphone (PSU); engineering plastics like liquid crystal polymer (LCP), polyetheretherketone (PEEK) and polyetherimide (PEI); and biodegradable materials such as polylactide. PMMA and PC are up to now the most popular polymer materials for microfabrication via hot embossing and injection molding. COC, a cyclo-olefin copolymer, is currently under test in the hot embossing process. This new material is extremely promising for applications in chemical engineering and molecular biotechnology since it has a high chemical stability and is optically transparent [17]. Tables 1 and 2 summarize the physical and chemical properties of the most commonly used thermoplastic polymers for micromolding. In comparison to glass, the resistance against chemicals, the aging, the mechanical stability, and the UV stability can restrict the use of polymers for certain applications. Also an increased fluorescence at shorter wavelengths (⬃400 nm, e.g., for dyes like fluoresceine) in comparison to glass can be ascertained, which can lead to a reduced sensitivity in case of laser-induced fluorescence (LIF) detection. Up to now there is no special development for plastics for the micromolding process since even a mass production of 1 million pieces leads to maybe 1 ton of polymer material—an amount much too small for special efforts. Therefore, already commercially available standard polymers are used for micromolding processes. However, the existing applications prove their principal suitability for microfabrication. For the fabrication of microfluidic devices, so far to our knowledge only thermoplastic and elastomeric materials have been used. 13.3 REPLICATION TECHNOLOGIES The secret of the commercial success of polymer microfabrication in microfluidics lies in the establishment of a low-cost manufacturing process. The underlying principle of these techniques is the replication of a microfabricated mold tool, which represents the negative (inverse) structure of the desired polymer structure. The (expensive) microfabrication step is therefore necessary only once for the fabrication of this master structure, which then can be replicated many times into the poly-

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Table 1

Basic Physical Properties of Molding Polymer Materials Heat distortion Linear

temperature

expansion

Measurement

Density

Glass

Permanent

Thermoplastic materials

(10 3(*)

temperature

temperature

conductivity

coefficient α

method:

for micromolding

kg/m 3 )

T g (°C)

of use (°C)

λ(W m ⫺1 K ⫺1 )

(10 ⫺6(*) K ⫺1 )

Vicat Method B (°C)

Thermal

Polyamid 6 (PA 6)

1.13

60

80–100

0.29

80

180

Polyamid 66 (PA 66) Polycarbonate (PC)

1.14 1.2

70 150

80–120 115–130

0.23 0.21

80 65

200 148–150

0.23–0.31

90–110

154–160

not available

60 (**)

Polyoxymethylene (POM)

1.41–1.42

Cycloolefin-copolymer (COC)

1.01 (**)

⫺60 138 (**)

90–110 not available

Data for the cycopentadien-

123 (**) Measurement

norbornen copolymer Zeonex

method: ASTM D648

Polymethylmetacrylate (PMMA)

1.18–1.19

106

82–98

0.186

70–90

80–110

Polyethylene low density (PE-LD)

ⱕ0.92

⫺10

70 (***)

0.349

140

40

Polyethylene high density (PE-

ⱕ0.954

—

90 (****)

0.465

200

60–65

0.22

100–200

90–100

HD) Polypropylene (PP)

0.896–0.915

Polystyrene (PS)

1.05

0–10 80–100

70 (*****)

0.18

70

78–99

Becker

(*) [19]. (**) Product information leaflet for Zeonex. (***) For Lupolen 1800a of BASF. (****) For Lupolen 6031 of BASF. (*****) For Polystyrol 159 K of BASF. Source: Ref. 18.

100

Basic Chemical Properties of Molding Polymer Materials

Thermoplastic materials for micromolding Polymid 6 (PA 6)

Polyamid 6,6 (PA 6,6)

Polycarbonate (PC)

Polyoxymethylene (POM)

Cycloolefin-copolymer (COC) (*)

Solvent resistance

Acid and alkaline resistance

Trade name

Resistant against: ethanol, benzine, aromatic and aliphatic hydrocarbons, mineral oils, fats, ether, ester, ketones same as above

Not resistant against: diluted and concentrated mineral acids, formic acid

Perlon Durethan (Bayer) Ultramid (BASF)

same as above

Resistant against: water, benzine, mineral oils Conditional resistant against: alcohols, ether, ester Resistant against: fuels, mineral oils, usual solvents Resistant against: acetone, methylethylketone, methanol, ethanol, isopropanol

Resistant against: diluted mineral acids

Nylon Nylind (Du Pont) Celanese (Ticona) Ultramid (BASF) Makrolon (Bayer)

Hostaform (Hoechst)

Topas (Ticona) Zeonex (Nippon Zeon)

325

Not resistant against: anorganic acids, acetic acid, oxidating solvents Resistant against: diluted and concentrated mineral acids and alcalines, 30% H 2 O 2 , 40% Formaldehyde, detergents in water

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Continued

Thermoplastic materials for micromolding Data for the cycopentadiennorbornen copolymer Zeonex Polymethylmethacrylate (PMMA) Polyethylene (PE)

Polypropylene (PP)

Polystyrene (PS)

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Table 2

Solvent resistance

Trade name

Not resistant against: concentrated HNO 3

Resistant against: up to 20% diluted acids, diluted alkalines, NH 3 Resistant against: NH 3, diluted HNO 3, H 2SO 4, HCL, KOH, NaOH Resistant against: most diluted acids and alkalines

Plexiglas (Ro¨hm) Lucryl (BASF) Perspex (ICI) Lupolen (BASF)

Resistant against: diluted and concentrated acids (except HNO 3) and alkalines

Polystyrol (BASF)

Hostalen (Hoechst)

Data for solvent, acid, and alkaline resistance taken from [18]. (*) Product information leaflet for Zeonex.

Becker

Not resistant against: ether, aromatic and aliphatic hydrocarbons, methylmethacrylate Resistant against: water, mineral oils, fuel, fatty oils Resistant against: alcohols, benzene, toluene, Xylene Resistant against: diluted solutions of salts, lubricating oils, chlorated hydrocarbons and alcohols Resistant against: alcohols, polar solvents. Not or hardly resistant against: ether, benzene, toluene, chlorated hydrocarbons, acetone, ethereal oils

Acid and alkaline resistance

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mer substrate. In addition to the cost advantages of this replication, it also offers the benefit of the freedom of design, as the master can be fabricated with a large number of different microfabrication technologies (see Sec. 13.4 below), which allow various geometries to be realized. Nevertheless, some restrictions apply to these replication techniques: 1. As the master has to be removed from the molded structure, undercuts (i.e., structures in the polymer with overhanging edges) cannot be fabricated by these means. 2. The primary success, the lifetime of the mold tool, and the achievable aspect ratios depend very strongly on the surface quality of the mold tool. In general, the smoother the tool surface, the lower the frictional forces on the tool as well as the polymer microstructure in the demolding step. Typically, surface roughness values of ⬎100 nm RMS are necessary to account for a good and reliable replication. This puts certain limitations on the fabrication methods for the mold tool. 3. The interface chemistry between tool material and substrate polymer also is a critical factor. If the two materials form any kind of chemical or physical bond during the replication step, this adds to the forces during the demolding step. Release agents, which are often used in the macro world to help the mold release of complex structures are often not suited for microfluidic devices, as on one hand they might diffuse into the polymer matrix and contaminate the sample if they are released into the liquid which is pumped through the channel; on the other hand, they tend to increase the autofluorescence of the polymer. With these criteria in mind, the following section describes the technologies available for the mold tool fabrication. 13.4 MASTER FABRICATION Various techniques have in the past been used for the fabrication of molding tools. We will concentrate on the methods relevant for microfluidic structures. 13.4.1

Micromachining Methods

As conventional machining has undergone miniaturization efforts as well, modern micromachining technologies (sawing, cutting, milling,

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turning) are capable of producing mold tools with structures down in the 10-µm range. Their particular advantage is the wide range of materials which can be machined, particularly stainless steel, which is not accessible with other microfabrication technologies and which offers very good mold insert lifetimes. In addition, the development times for micromachined tools can be shorter, as no mask fabrication and lithography step is involved. Especially comparatively simple channel structures with straight walls are well-suited geometries for these techniques. Channel crossings, high aspect ratio structures, very deep holes, or very small structures, however, cannot or only with major drawbacks be fabricated with these methods. Special mention here should be made of microelectrodischarge machining (µ-EDM), which allows the fabrication of quasi three-dimensional structures in conducting materials [20]. In this technique, the material is removed due to the high-energy electric discharge between an electrode and the workpiece. It offers a high degree of flexibility in terms of materials and geometries, but produces a comparatively rough surface. This method nevertheless promises a good potential for further development. Very simple structures have reportedly been fabricated by replicating thin chromel wires [21,22]. 13.4.2

Electroplating Methods

The most commonly used methods for master fabrication employ an electroplating step, resulting in a replication master made out of nickel or a nickel alloy like NiCo or NiFe. The process starts with a photolithography step, where a substrate with a conducting electroplating starting layer which is coated with photoresist is exposed to light. The resist is subsequently developed, so that the areas which should be electroplated are free of resist. This structure is then placed into a galvanic bath, where due to the migration of metal ions between the bath and the conducting starting layer, the metal starts to grow in the resist structure. After the resist structure is overgrown with the metal, the resist and starting layer can be dissolved and the resulting metal structure (so-called shim) can be processed further. Conventional photoresist technologies allow structural heights of the order of 10–40 µm, for higher structures special thick resists like EPON SU-8 [23] have to be

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used, which can result in heights of up to one mm. Other techniques to produce electroplating forms are the LIGA technique (German acronym for Lithographic (lithography), Galvanoformung (electroplating), Abformung (molding)), where thick PMMA layers are exposed with synchrotron radiation [24] and the laser-LIGA process [25], where the synchrotron radiation is replaced by pulsed UV light, which ablates the polymer material. These techniques have in common that the surface roughness is quite small (LIGA down to ⬃10 nm RMS) and that the resulting nickel tool has a good surface chemistry for most polymers. Drawbacks are (1) the comparatively slow growth rate of nickel in the electroplating process (a typical rate is typically between 10 and 100 µm/hr); (2) the high stress levels in thick nickel layers which tend to bend the master; and (3) the radial dependency of the growth rate which can result in a different height of the nickel structure in the middle and at the rim of a nickel wafer. In addition the cost structure of LIGA seems to handicap its commercial breakthrough. 13.4.3

Silicon Micromachining

As silicon itself has suitable material properties for a mold tool (high stiffness, high heat conductivity) and a large variety of silicon surface micromachining techniques exist as well as commercially available services, attempts have been made to use silicon directly as a tool material. Several micromachining technologies have been under investigation, the simplest one being wet etching of silicon. A wet etching step of 100-silicon results in a structure with a wall angle of 54.7°, which forms a trapezoidal channel. The slant in the wall allows a good mold release, and the surface roughness of the wet etching process of well-oriented monocrystalline silicon wafers is excellent [21,22,26,27]. Obviously, the channel cross section in this case is limited to this shape, although some isotropic etching techniques for silicon also exist. This limits the achievable aspect ratio. Dry-etching methods (reactive ion etching, RIE, advanced silicon etch, ASE, or the Bosch process; for a review see e.g. [16]), however, allow deep structures with vertical walls to be fabricated, alas with a surface roughness which strongly depends on the etch rate. Fast etches produce a very rough surface, which for practical reasons limits the

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achievable depth of the structures. Typical depth range between 10 and 40 µm for a conventional dry etch and up to ⬎200 µm in case of an ASE process, with etch rates of the order of 1–5 µm/min. While in the past tool fabrication with advanced silicon etch processes suffered from this high roughness of the side walls and undercuts at the bottom of etched trenches, novel process developments [28] show great potential for the use of dry-etched silicon structures as embossing tools. All silicon tools, however, have the common problem of a potential stiction problem with many polymers due to their surface chemistry. A combination of silicon etching and electroplating exists, which is called DEEMO (deep etching, electroplating, molding) [29], where the micromachined silicon is used as a base for an electroplating step equivalent to the electroplating of the photoresist described above.

13.5 HOT EMBOSSING PROCESS Currently the most widely used replication process to fabricate channel structures for microfluidic applications is hot embossing [21,22,27,30– 36]. The microfabrication process of hot embossing is itself a rather straightforward one [37]. The process is shown schematically in Figure 1. After obtaining the embossing master, it is mounted in the embossing

Figure 1 Diagram of the hot embossing process.

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machine (see Fig. 2). An embossing machine typically consists of a force frame which delivers the embossing force via a spindle or a hydraulic system and a T-bar to the boss. The embossing tool and the planar polymer substrate are mounted on heating plates, which can also contain cooling channels. In these channels, a high-heat-capacity oil is circulated in the cooling phase which allows active cooling with cooling times equivalent to heating times at ⬃1 min between upper and lower cycle temperatures. This configuration allows an isothermal heating and cooling of both tool and substrate. At the beginning of the embossing cycle, the two are heated separately in a vacuum chamber at about 10 ⫺1 mbar to a temperature just above the glass transition temperature T g of the polymer material. The vacuum, although extending the overall cycle time due to pumping and venting times, is necessary for a good-quality replication. In addition it prevents nickel tools from oxidation at elevated temperatures, which extends the lifetime of the mold. For most standard thermoplastic materials like polymethylmetacrylate (PMMA) or polycarbonate (PC), this temperature is in the range of 100–180°C (PMMA 106°C, PC 150°C). The tool is brought into contact with the substrate and then embossed with a sensor feedback controlled force, typically of the order of 20–30 kN for a 4″ embossing tool. Still applying the embossing force, the tool-substrate sandwich is then cooled to just below T g to stabilize again the polymer microstructure. To minimize thermally induced stresses in the material as well as replication errors due to the different thermal expansion coefficients of tool and substrate (typical values for the thermal expansion coefficient of polymers are of the order of 7 ⫻ 10 ⫺5 /K, while silicon has ⬃1.5 ⫻ 10 ⫺6 /K and nickel around 1 ⫻ 10 ⫺5 /K), this thermal cycle should be as small as possible, in our case currently about 30°C. After reaching the lower cycle temperature, the embossing tool is mechanically driven apart from the substrate which now contains the desired features. This is usually the most critical step, as now the highest forces act on the polymer microstructure, particularly if a structure is desired with vertical walls and a high aspect ratio. Therefore, an automated mold release is crucial for a high production yield. Overall cycle time of the embossing process for materials such as PMMA is of the order of 5–7 min.

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(a)

(b)

Figure 2 (a) Schematic of a hot embossing machine. (b) Hot embossing machine HEX 03.

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Table 3 gives an overview for the process parameters used for the embossing of PMMA and PC. For every design of a microstructure, the process conditions vary slightly, as properties of the design like the distribution of large and small structures over the wafer area, total processed wafer area and geometry of the structures (i.e., radius of curvature, free-standing structures or connected structures) come into play. Therefore the parameters given in Table 3 vary in the range indicated. The term ‘‘Hold Time’’ denotes the time at the upper cycle temperature with applied embossing force. The main difference in this process in comparison to existing replication techniques like in the CD industry lies in the fact that both the tool and the substrate are thermocycled here, while for CD manufacturing, the heated material is molded in a cool cavity which remains at a constant temperature below T g. To achieve higher aspect ratios than on a CD (aspect ratio of CD pits ⬃0.2) and a good mold fill, this thermocycling however is necessary. To emboss high aspect ratio structures, in addition the following process and material properties have to be taken into account: 1. Sidewall roughness of the embossing master. To obtain an undamaged structure, the frictional forces between the embossing tool and the polymer microstructures in the deembossing process have to be minimized. The microstructure is destroyed if the frictional forces become larger than the local tensile strength of the polymer. Therefore in the master fabrication process care has to be taken to insure a minimal sidewall roughness; 80 nm RMS is an empirical limit for the fabrication of structures with an aspect ratio ⬎0.5. For this reason, the LIGA process is perfectly suited for the high aspect ratio structure fabrication as the side wall roughness can be as little as 10 nm RMS. Recently, a surface roughness of only 8 nm has been reported for ASE processed silicon [28], which would allow for very good replication properties. 2. Sidewall angles. The above-described friction between master and substrate is particularly critical in microstructures with vertical sidewalls. Already a small deviation from the vertical eases this constraint, as mechanical contact between tool and substrate is immediately lost in the deembossing step. If the application and master fabrication allow wall angles different from 90° [38], this is advantageous in the fabrication process.

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Material and Process Conditions for the Embossing of PMMA and PC

Material

Density (10 3 kg/m 3 )

PMMA PC

1.17–1.20 1.20

T g (°C)

Young’s modulus (MPa)

Embossing temperature (°C)

Deembossing temperature (°C)

Embossing force (4″) (kN)

Hold time (sec)

106 150

3100–3300 2000–2400

120–130 160–175

95 135

20–30 20–30

30–60 30–60

Process data range represents the dependence on structural design parameters.

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3. The chemical interface between master and substrate. To minimize stiction between embossing master and the polymer material, which creates an additional force in the deembossing step, both surfaces should offer as little chemical surface bonding sites as possible. Although release agents can be added to the polymer which allow the realization of aspect ratios of up to 50 [39], applications in the life sciences prohibit the use of these materials due to a potential sample contamination. 4. Temperature coefficients. As in the hot embossing, a temperature gradient is involved, the difference in the temperature coefficients of tool and substrate material has to be taken into account to avoid the creation of additional forces due to the larger shrinkage of the polymer material in comparison to the tool material. For any commercial production of microstructured components, a high throughput of the fabrication technology is crucial. In the replication methods this translates to a multiple tool or to an as large area for embossing as possible. We have generally utilized a 4″ embossing area to fabricate our polymer microstructures, which requires embossing forces of the order of up to 30 kN. 13.6 EXAMPLES Figures 3 and 4 show the injection region of a capillary electrophoresis chip. The injection volume is defined by the channel intersection, so the geometrical accuracy of the replication process is of importance. Figure 3 shows the injection region embossed in PMMA with a LIGA fabricated nickel tool, which yields vertical side walls. In Figure 4, the embossing master was a wet etched silicon wafer, therefore the walls display the typical 54.7° wall angle. Such a tool has the advantage that in the mold release step the physical contact between embossing tool and structure is immediately lost, and therefore structural deformation due to frictional or shear forces between the embossing master and the substrate is avoided. The accuracy of the replication process can be assessed in the inset, where etching defects of the tool due to a crystalline misorientation are clearly replicated. The estimated step height is ⬍100 nm. For higher aspect ratios and for structures with vertical walls, an

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Figure 3 Injection region of a capillary electrophoresis chip fabricated with a nickel tool in a PMMA substrate.

advanced silicon etch process (ASE) in an ICP reactor has been used. The design of the structure, a two-dimensional microchannel array for capillary electrophoresis applications like protein analysis, was previously reported elsewhere [13]. To achieve a maximum separation efficiency, very small channels with a high aspect ratio had to be fabricated. A cross section of the silicon tool can be seen in Figure 5a, the ridges being 0.8 µm wide, 5 µm high, with a 5 µm pitch and with a total of 500 channels. In Figure 5b this tool has been replicated in PMMA, the submicron channel array with an aspect ratio of about 5 can be clearly seen, as well as the slight structural distortion due to the mold release. In Figure 6, a separation of 10 ng/mL HaeIII ΦX174 DNA fragments in 0.1X Tris-borate-EDTA buffer (TBE) is shown. Although not optimized for separation speed, this result clearly shows the advantages of minaturization in terms of analytical speed. Other examples of microfluidic structures are given in Fig. 7 and 8. Figure 7 shows a structure for fluid mixing with channels of a cross section of 50 ⫻ 50 µm in a polycarbonate substrate (structural design: O. Geschke, MIC, Lyngby, (text continues on p. 341)

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Figure 4 Injection region of a capillary electrophoresis chip fabricated with a wet-etched silicon tool in a PMMA substrate.

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Figure 5 (a) Embossing tool fabricated with an advanced silicon etch; (b) resulting channel array structure.

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Figure 6 Separation of HaeIII ΦX174 DNA fragments in a hot-embossed PMMA chip. Data gratefully supplied by A. Paulus (Acalara BioSciences).

Figure 7 Fluid mixing chip in PC (Design O. Geschke, MIC, Lyngby).

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Figure 8 (a) Silicon tool for a frit/filter. (b) Replicated structure in PMMA.

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Denmark). Figure 8a shows the etched silicon tool for a frit/filter structure, and Figure 8b the structure replicated in PMMA. Other typical application examples are nanowell plates [38] with well diameters varying from 500 to 25 µm.

13.7 ADDITIONAL MANUFACTURING TECHNOLOGIES Besides the process of generating microchannels or microchannel networks, several other manufacturing steps have to be addressed to account for a completed microfluidic device. This includes most importantly the closing of the microchannels to form capillaries, but also refers to the dicing of devices, via hole fabrication and possible inclusion of metal structures. All these steps are necessary to create a fully functional microfluidic system. 13.7.1

Bonding

After the microfabrication process, the microchannels in the polymer substrate are still open. They have to be closed to form true capillaries. Potential fabrication problems in this process are the clogging of the channels, a change in their physical parameters, and the alteration in their dimensions. This often represents a big challenge for highervolume fabrication methods. Several methods have been reported in the literature: Lamination This method is widespread in the macroworld for encapsulation of paper and polymers in a polymer film and works well for comparatively large channels. In the lamination process, a thin PET foil (typical thickness ⬃30 µm) coated with a melting adhesive layer (typical thickness 5–10 µm) is rolled onto the structure with a heated roller [40–43]. The adhesive layer melts in this process and combines the lid foil with the channel plate. With very small channels, however, the adhesive tends to block the channel. Due to the difference in the materials used, a nonhomogeneous interface between lid and channel plate is created which leads to a sudden change of parameters such as refractive index

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at the interface. Figure 9 shows the cross section of such a microchannel, closed with a PET/PE laminate [41]. Gluing Similar to lamination, conventional gluing can be used to join channel plate and lid [42,44], whereby the same problems arise, mainly the risk of blocking the channels. Application of Heat and Pressure Several teams report a sealing of structures by heating up the polymer and applying a force to close the channels [40]. Care has to be taken not to damage the structures, so this method is advisable mainly for designs with comparatively small structured areas in comparison to the whole chip surface. Laser Welding Polymers can be joined by local melting due to heat generated by a laser. This has been successfully demonstrated in the fabrication of micropumps [45], but so far no reports have been published on microchannel applications. The main reason for this is that all welding lines have to be drawn out with the laser, which in case of microchannels amounts to comparatively large distances and therefore welding times. Novel beam-shaping setups, which illuminate large areas through a mask, can overcome this problem [46]. However, the requirement of at least one laser light-absorbing (i.e., opaque) polymer sheet remains, so that optical observation of the channel is restricted to one side of the chip. Ultrasonic Welding A method well known in the macroworld is ultrasonic fusion of two polymer layers, where a local melting of the polymer is achieved by the energy density of an ultrasonic sound wave. To date, to our knowledge no application of this technique has been published in microfluidics. Common for all these bonding technologies is the need for very clean processing conditions, as especially particle contamination re-

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Figure 9 Laser-ablated microchannel laminated with a PET/PE foil. (From Ref. 41.)

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Figure 10 Laser-cut PMMA lid on top of microchannels.

duces the bond quality and yield dramatically. Clean room processing is therefore highly advised. 13.7.2

Dicing and Hole Drilling

Dicing the devices can be realized either with a conventional rotating saw (no wafer saw; the polymer tends to smear the cutting wire) or with lasers, mainly CO 2 lasers with little energy (typically 50 watts). The method of choice depends on the material used and the accuracy required. Figure 10 shows a cover lid on top of a microchannel system. The surface topology is mainly given by the melting characteristics of the polymer, as the infrared laser light melts the material along the cut line (in contrast to UV laser light, which is used in laser ablation, where the polymer chains are broken by the UV radiation and the reaction products evaporate). The surface roughness is on the order of 10 µm. Holes can also be fabricated either conventionally with a mechanical drill or with laser drilling. An example for the achievable quality of mechanically drilled holes (diameter 1 mm) can be seen in Figure 11. These holes act as fluidic reservoirs and interfaces to the macroscopic world in a microfluidic chip. The wall roughness is ⬎5 µm, but the more critical

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Figure 11 Mechanically drilled hole in PMMA. Note the undistorted microchannel branching off into the chip at the bottom of the well.

parameter is the deformation of the hole edge (burr). Any burr formation larger than a few micrometers leads to an incomplete bonding at that location. Therefore the drilling procedure has to be optimized in terms of drill speed and applied force. Nevertheless, this figure shows that it is possible even with ordinary ‘‘macroscopic’’ drills to achieve the necessary hole and edge quality. A microchannel with a width of 50 µm and a height of 25 µm can be seen to branch off the drilled reservoir into the chip, and a cover plate has been bonded to the hole plate to seal the channel (bottom of the picture). 13.7.3

Electrode Fabrication

If electrode structures for amperometric detection [47,48] or the application of the separation voltage should be included, the normal deposition methods such as sputtering and thermal or electron beam evaporation can be used. A limitation exists in the achievable dimensions of the electrode structures, as the deposition can easily only be done with a shadow mask, which restricts the electrode width to ⬃40 µm and above. Figure 12 shows an example of a Ag reference electrode on top of a microchannel. Photolithographic processes are difficult to carry out on an already structured polymer chips.

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Figure 12 Metal electrode deposited over a microchannel.

13.8 OUTLOOK AND CONCLUSIONS As more and more commercial applications for microfluidic devices come into existence, polymer microfabrication methods are becoming increasingly important. These application areas all demand a high number of devices at low cost. Most of the devices will be used as disposables. This field is still in its infancy, however, with many fabrication and material issues still unaddressed. Nevertheless an enormous growth in publications can be observed. Nowadays, hardly any conference in the field of µ-TAS lacks a session on polymeric devices, which is in stark contrast to the situation only 5 years ago, when at µ-TAS only a single paper addressed this issue. Besides the commercialization of microfluidics with its applications in genomics, drug discovery, and diagnostics as a driving force for the development, we can observe that µ-TAS emancipates itself from its historical fabrication roots which came from microelectronics and develops fabrication technologies which are better suited for the above mentioned applications in the life sciences. What are the main trends in this exciting field? First, several groups work in the integration of either sample preparation issues and/ or detection systems onto the fluidic chips. This is a development to-

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wards a ‘‘real’’ miniaturized system. Second, the first applications with a demand for a very high number of devices (several million a year) emerge, which certainly will stimulate development in the fabrication technologies. Third, more attention will be directed into the materials themselves, as the range of polymeric material is much wider than the scope currently under investigation; the tuning of material parameters for specific applications will become an important topic, as well as potential surface modifications. We have just seen the beginning of these developments, but the commercialization will prove a tremendous stimulus. In addition, more and more academic groups realize the great potential for simple and fast in-house production of design prototypes with polymer fabrication methods. Polymer based systems will certainly become ‘‘household’’ items in the years to come.

ACKNOWLEDGMENTS The author would like to thank Claudia Ga¨rtner from amt Jena for the help in preparing the manuscript and Aran Paulus from Aclara BioSciences for the separation data. The work on the 2D-CE chips was carried out in the group of Professor Andreas Manz at the Department of Chemistry, Imperial College, London, and at IMM Mainz under EU contracts CHGE-CT93-0052

REFERENCES 1. Manz, A., Becker, H., eds. Microsystem Technology in Chemistry and Life Science, Topics in Current Chemistry 194. Springer, Heidelberg, 1998. 2. http://www.the-scientist.library.upenn.edu/yr1997/sept/ latta_p1_970915.html 3. Terry, S.C., Jermann, J.H., Angell, J.B. IEEE Trans. Electron. Devices ED-26 (1979), 1880–1886. 4. Manz, A., Graber, N., Widmer, H.M. Sensors Actuators B 1, (1990), 244–248. 5. Manz, A., Fettinger, J.C., Verpoorte, E., Lu¨di, H., Widmer, H.M., Harrison, D.J. Trends Anal. Chem., 1991, 10(5), 144–149.

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6. Harrison, D.J., Fluri, K., Seiler, K., Fan, Z., Effenhauser C., Manz, A. Science, 1993, 261, 895–897. 7. Fan, Z., Harrison, D.J. Anal. Chem., 1994, 66(1), 177–184. 8. Effenhauser, C.S., Manz, A., Widmer, H.M. Anal. Chem., 1993, 65(19), 2637–2642. 9. Jacobson, S.C., Hergenro¨der, R., Koutny, L.B., Ramsey, J.M. Anal. Chem., 1994, 66(14), 2369–2373. 10. Jacobson, S.C., Hergenro¨der, R., Moore, J.A.W., Ramsey, J.M. Anal. Chem., 1994, 66(23), 4127–4132. 11. Jacobson, S.C., Moore, A.W., Ramsey, J.M. Anal. Chem., 1995, 67(13), 2059–2063. 12. Jacobson, S.C., Ramsey, J.M. Electrophoresis, 1995, 16, 481–486. 13. Becker, H., Lowack, K., Manz, A. J. Micromech. Microeng., 1998, 8, 24–28. 14. Becker, H., Manz, A. Integrated capillary electrophoresis for chemical analysis. In: Baltes, H., Go¨pel, W., Hesse, J. eds. Sensors Update, Vol. 3. VCH, Weinheim, 1998, 208–238. 15. Kopp, M.U., Crabtree, H.J., Manz, A. Curr. Opin. Chem. Biol. 1 (1997), 410–419. 16. Jansen, H.V., Gardeniers, J.G.E., de Boer, M.J., Elwenspoek, M.C., Fluitman J.H.J. J. Micromech. Microeng., 1996, 14–28. 17. Ehrfeld, W., Hessel, V., Lo¨we, H., Schulz, Ch., Weber, L. Microsystem Technol. 5 (1999) 105–112. 18. Taschenbuch der Werkstoffe, Merkel, Thomas. Fachbuchverlag, Ko¨ln, 1994. 19. CRC Handbook of Chemistry and Physics, 73rd ed. CRC Press, Boca Raton, 1993. 20. Weck, M., Fischer, S., Vos, M. Nanotechnology 8 (1997), 145–148. 21. Martynova, L., Locasico, L.E., Gaitan, M., Kramer, G.W., Christensen, R.G., MacCrehan, W.A. Anal. Chem. 69 (1997), 4783–4789. 22. Locascio, L.E., Gaitan, M., Hong, J., Eldefrawi, M., Proc. Micro-TAS ’98. Banff, 1998, 367–370. 23. Despont, M., Lorenz, H., Fahrni, N., Brugger, J., Renaud, P., Vettiger, P. Proc. MEMS’97, Nagoya, 1997, 518–523. 24. Ehrfeld, W., Mu¨nchmeyer, D. Nucl. Instrum. Methods A303 (1991), 523–532. 25. Arnold, J., Dasbach, U., Ehrfeld, W., Hesch, K., Lo¨we, H. Appl. Surf. Sci. 86 (1995), 251–258.

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26. McCormick, R.M., Nelson, R.J., Alonso-Amigo, M.G., Benvegnu, D.J., Hooper, H.H. Anal. Chem. 69 (1997), 2626–2630. 27. Becker, H., Heim, U. Sensors Materials 11, (1999), 297–304. 28. Chabloz, M., Sakai, Y., Matsuura, T., Tsutsumi, K. Proc. HARMST’99. Kisarazu, 1999, 26–27. 29. Elders, J., Jansen, H.V., Elwenspoek, M., Ehrfeld, W. Proc. MEMS ’95. Amsterdam, 1995, 238–244. 30. Niggemann, M., Ehrfeld, W., Weber, L. Proc. SPIE Micromachining and Microfabrication Process Technology IV, Vol. 3511. Santa Clara, 1998. 31. Becker, H., Dietz, W., Dannberg, P. Proceedings Micro-TAS ’98. Banff, Canada, 253–256. 32. Becker, H., Dietz, W. Proc. SPIE Micro Fluidic Devices and Systems. Santa Clara, 1998, 177–182. 33. Konrad, R., Ehrfeld, W., Hartmann, H.J., Jacob, P., Pommersheim, R., Sommer, I. Proc. 3rd Intl. Conference on Microreaction Technologies, 18.–21.4. 1999, Frankfurt, 420–429. 34. Becker, H., Heim, U., Ro¨tting, O. Proc. SPIE ‘‘Microfluidic Devices and Systems II’’, Vol. 3877, Santa Clara 1999, 74–79. 35. Becker, H., Heim, U., Ro¨tting, O. Proc. Micro-Engineering ’99, Stuttgart, 1999, 74–80. 36. Bachmann, M., Chiang, Y.M., Chu, C., Li, G.P. Proc. SPIE Microfluidic Devices and Systems II, Vol. 3877. Santa Clara, 1999, 139–146. 37. Heckele, M., Bacher, W., Mu¨ller, K.D. Microsystem Technol. 4 (1998), 122–124. 38. Becker, H., Klotzbu¨cher, T., Proc. 3rd Int. Conf. on Microreaction Technologies, 18.–21.4. 1999, Frankfurt, 102–112. 39. Ro¨tting, O., Ko¨hler, B., Reuther, F., Blum, H., Bacher, W. Proc. SPIE Design, Test and Microfabrication of MEMS/MOEMS, Vol. 3680(2). Paris, 1999, 1038–1045. 40. Paulus, A., Williams, S.J., Sassi, A.P., Kao, P.K., Tan, H., Hooper, H.H. Proc. SPIE Microfluidic Devices and Systems, Vol 3515. Santa Clara, 1998, 94–103. 41. Roberts, M.A., Rossier, J.S., Bercier, P., Girault, H., Anal. Chem. 69(11) (1997), 2035–2042. 42. Soane, D.S., Soane, Z.M., Hooper, H.H., Alonso-Amigo, M.G. Int. patent Appl. WO 98/45693, 1998. 43. Schwarz, A., Rossier, J.S., Bianchi, F., Reymond, F., Ferrigno, R., Girault, H.H. Proc. Micro-TAS’98. Banff, 1998, 241–244.

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¨ hman, O., Sjo¨din, H. Int. patent WO 91/ 44. Ekstro¨m, B., Jacobsen G., O 16966, 1990. 45. Ka¨mper, K.P., Do¨pper, J., Ehrfeld W., Oberbeck, S. Proc. MEMS’98. Heidelberg, 1998, 432–437. 46. Chen, J.W., Hessler, T. Proc. Micro-Engineering ’99. Stuttgart, 1999, 81–88. 47. Woolley, A.T., Lao, K., Glazer, A.N., Mathies, R.A. Anal. Chem. 1998, 70, 684–688. 48. Rossier, J.S., Roberts, M.A., Ferrigno, R., Girault, H.H. Anal. Chem. 1999, 71, 4173–4177.

14 Microfabricated Reactor Technology Tibor Chova´n University of Veszpre´m, Veszpre´m, Hungary

Andra´s Guttman Torrey Mesa Research Institute, San Diego, California

14.1 INTRODUCTION Adapted from the microelectronics industry, the notion of miniaturization was applied to chemical and biochemical engineering. Microfabricated chemical and biochemical reactor technology is anticipated to offer numerous advantages in chemical processing and biotechnology production [1,2]. The improved heat and mass transfer properties typical of microfabricated systems enable the use of more intensive reaction conditions also resulting in higher yields than that of conventional size reactors. On the other hand, the increased risk and capital cost associated with scaling up from laboratory to production size limit the introduction of new and/or expensive reagents. However, scale-up to production by parallelization of microreactor units used in the development process reduces the cost of redesign and pilot plant experiments, hence, yield rapid development from laboratory to commercial production scale. The laminar flow profile in the reduced dimensions of microdevices enables accurate heat and mass transfer characterization, as well as easy extraction of kinetic parameters from the available data. Spatial and temperature control of reagents and reactants can be 351

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obtained under diffusion-limited and unique thermal properties that exist at micrometer scale [3]. Miniaturization, in conjunction with integration of multiple functionalities, has the capability to construct structures that exceed the performance of traditional macroscopic systems and also provide a plethora of new functionalities and the potential of low-cost mass production [4]. Sample preparation, purification, mixing, reactions, separations, and fraction collection all can be performed on a highly integrated monolithic microfabricated device. Applicability of recently developed microfabrication reactor technologies commenced a rapidly moving interdisciplinary area of miniaturization of chemical and biotechnological processes. Since the introduction of photolithographic techniques for the fabrication of chemical and biochemical microdevices [5] the number of applications has been exponentially increased [6]. 14.2 MICROFABRICATED REACTORS The especially large surface-to-volume ratio is one of the most important properties of microreactors [7]. Not only can extremely rapid and highly exothermic reactions be accomplished under isothermal conditions, but also the mass transfer distances are very low in the case of multiphase reactions. Thus, higher selectivities are expected and more precise kinetic information can be obtained, that would not be available otherwise in conventional scale methods [8]. Microreactors make feasible the realization of on site production at the point of demand. The excellent heat transfer characteristics of microfabricated devices also avoid the risk of potentially significant industrial accidents caused by thermal runaway [9]. 14.2.1

Microfabricated Gas-Phase Reactors

Microreactors for heterogeneous gas-phase reactions are typically constructed in the form of microchannels. Catalytic microreactors are constructed by depositing the catalyst on the inner microchannel surface of micromixers and heat exchangers. In case of highly exothermic reactions, heat exchange plates can be inserted into the microchannel structure. The selectivity which has critical influence on the reactor perfor-

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mance of many catalytic reactions, like partial oxidation, is readily controlled by regulating catalyst temperature. A heterogeneously catalyzed gas-phase reaction, oxidative dehydrogenation of an alcohol to aldehyde, was studied by Wolf and coworkers [8] to understand a scaleup problem that occurred during process development. In a microreactor, using silver as catalytically active material, they could establish a short enough residence time getting in this way an adequately high heat transfer rate compared to a multi-short-tubular reactor and got an appropriate development tool. Operating a continuous reactor in unsteady state or periodic regime can yield significantly higher selectivity and reactor performance than that of optimal steady-state operation [10]. The optimal cycle time of periodic operation may range from fractions of a second up to hours. Note that microfabricated reactors are more appropriate for periodic operation at high frequencies than conventional macroscopic reactors. Owing to the quite small radial diffusion times, the residence time distribution (RTD) is very narrow in the microchannels. Renken and coworkers [11] studied the catalytic dehydration of isopropanol using periodic operation mode. The catalyst was a washcoated γ-Al 2O 3 layer in the microchannels of a stainless-steel reactor. The behavior of the microreactor, constructed of stacked plates, was similar to that of a cocurrent or countercurrent heat exchanger. The reactor design was based on the structure of earlier reported micro-heat exchangers [12]. 14.2.2

Microfabricated Liquid-Phase Reactors

Providing adequate mixing is one of the most important fundamental design and operation problems of liquid-phase microreactors. Owing to the small dimensions of microfabricated devices, Reynolds numbers are low and the flow profile is generally laminar; consequently, mixing mainly occurs by diffusion [4]. The fact that multiple streams are mixed very slowly enable exploiting further possibilities for phase transfer reactions and separation devices [13,14] and can be utilized in novel microfabrication schemes [15]. In order to decrease diffusion lengths and increase the interfacial area, mixing is usually performed by repeated lamination of the streams to be mixed [16]. One method to obtain rapid diffusive mixing is hydrodynamically focusing the side flows

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into the inlet flow as a narrow stream [17]. First the static mixer setup contacts two fluid streams, then it divides the resulting stream into two flows perpendicular to the interface area. The following recombination of these two streams then results in doubling of the mixing interface and halving of the diffusion length. The process is repeated several times and ultimately provides rapid, intensive mixing due to the significantly shorter diffusion paths [18,19]. A reaction catalyzed by concentrated sulfuric acid forming a second phase was studied by Worz et al. [8]. The reactant was dissolved in hexane and the product was transferred into the acidic phase. Quick side reactions of the reactant, the intermediate, and the product led to formation of byproducts. A two-stage reactor scheme was applied. The first reactor of 32 reaction channels had very short residence time (1 sec) while the second reactor provided residence times in the minute range. This approach allowed higher selectivity and yield. A relevant structural element of this liquid-liquid microreactor is shown in Figure 1. When the reaction rate does not exceed the rate of micromixing, liquid-phase screening is possible with no extra difficulty. On the other hand, processes such as hydrogenation, carbonylation, and hydrofor-

Figure 1 Structural element of a multichannel, liquid-phase microreactor. (From Ref. 8.)

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mylation work in multiphase systems (i.e., gas/liquid or gas/liquid/ liquid phases). Inappropriate phase and catalyst manipulation originating from inadequate mixing may drastically influence selectivity and reactivity estimations. A test reactor designed for high-throughput screening using low amounts of catalysts with sufficiently long residence times were tested on liquid/liquid isomerization and gas/liquid asymmetric hydrogenation reactions [20]. The approach applied continuous carrier flows, and simultaneously injected substrate and catalyst pulses to a micromixer with short residence times followed by application of a tubular reactor with long residence time. Some of the advantages of this method over traditional parallel-batch operations are the reduced sample amounts (to microgram levels), the increased range of operating conditions (pressure, temperature), and the fewer and simpler electromechanical moving parts. The scheme of the high-throughput microreactor is presented in Figure 2. A prototype of a continuous flow, combinatorial screening microreactor, which can be used for rapid synthesis of a line of reaction products, was developed by Wong and coworkers [21]. Their microdevice employed electro-osmotic (EOF) and electrophoretic flows as driving force for reagent mobilization. Applying electro-osmotic flow instead of pressure-driven flow has apparent advantages, such as experimental simplicity and capability to maintain reagent flow without any moving parts while the back-pressure effects are minimal. A further advantage is that electric field-mediated separation can be achieved as well. An ideal solution is envisioned as a new generation of microfabri-

Figure 2 High-throughput sequential screening reactor. S, substrate; P, product; cat, catalyst. (From Ref. 20.)

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cated devices, which integrates the screening microreactors with the already existing µTAS separation devices. 14.2.3

Microfabricated Packed-Bed Reactors

It is not obvious to elucidate the results, obtained by using thin-layer catalysts, to the porous catalyst support applicable in the industry. This emphasizes the importance of the development of microreactors employing porous support catalysts manufactured by conventional procedures. Microreactors represent an obvious choice for combinatorial and statistically planned applications; however, only a limited number of catalysts can be used in thin-layer format. The high surface-to-volume ratios characteristic of microfabricated structures result in improved thermal and mass transfer and suggest that microfabricated multiphase systems could have performance advantages relative to conventional macroscopic systems, or at least small-scale productions. Microfabrication also provides unique opportunities to create catalyst support systems, which avoid packing variations associated with distribution of catalyst particle sizes [22]. The size of conventional reactors can be significantly decreased without any loss in the expected throughput by using microchannel formats. An example is the application of fuel cells as mobile or stationary power sources relying on a fuel processor demanding highly sophisticated reactor technology. The fuel processor can convert hydrocarbon-based raw materials to high hydrogen content products in a few processing steps including the water gas shift (WGS) reaction. Wegeng’s group [23] experimentally studied the WGS reaction on engineered monolithic catalyst in a planar-sheet-architecture microreactor. They showed very fast kinetics of the WGS reaction (millisecond scale), consequently proving that small (10–100 times smaller than the conventional reactors), modular, high-performance, low-cost microreactors can be designed for fuel processing purposes. Recently, Harrison and coworkers [24] described a novel design, utilizing microweirs to reversibly trap reagent-carrying beads within a microchannel structure, allowing solutions to pass through the packed bed. Packing the chamber was accomplished by electrokinetic or pressure-mediated bead filling through an introduction channel into the space formed by

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the two weirs. The ability to pack and retain beads within the microchip was utilized to develop bead-based on-the-chip solid-phase extraction (SPE), electrochromatography (EC), and immunoassays. 14.2.4

Unit Operations

Unit operation development involves building and testing the microdevice at the actual size. The use of microfabricated elements can potentially intensify the processing capability of any unit operations in chemical and biochemical processing. Advantages are expected by miniaturization are lightweight and compact system design, superior heat and mass transport, precisely controllable operation parameters, high-throughput per unit volume, cost economies at mass production, and feasible distributed and mobile applications. Should the throughput be increased, further units with the same parameters are combined in a parallel fashion. This also implies that during product development the time frames for the setup, testing, and turnaround are much smaller than in the case of traditional technology development [25]. Different types of micromixer units with various operating principles and parameters have been designed and applied effectively in numerous applications, like emulsion and foam formation, multiphase reactions, and polymerization. For gas-liquid processes a device called a microbubble column, based on a static micromixer with an interdigital channel structure, can provide dispersing of gas bubbles into the liquid phase [26]. Due to the anticipated difficulties of inducing electrophoresis and electro-osmosis in organic solvents, it is essential that for both aqueous solutions and organic solvents a pressure-driven flow can be applied as means of fluid transportation. Kitamori’s group [27] studied a solvent extraction process using a microfabricated device. Two liquid streams entered into a microchannel and produced in this way a two-phase laminar flow. When continuous flow was applied, the two phases did not mix with each other, and a stable liquid-liquid (aqueous-organic) interface evolved in the microchannel. A general approach to the application of Micro-Chemical and Thermal Systems (Micro-CATs) can be based on microcomponent unit operations, like micromixers, micro heat exchangers, micro gas absorbers and absorbers, microchannel catalytic reactors, micro-

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channel electrochemical reactors, microchannel gas/liquid separators, and other microreactors and separators. These operations can provide high levels of temperature and consequent reaction control, as well as thermal integration and parallel operation [28]. The miniaturization of complex chemical processes is possible by applying the recent developments of microtechnology for all conventional chemical engineering unit operations including heat exchangers [29], reactors [30], separators [31], and actuators [32]. An integrated microreaction system combining mixing and heat exchanger units is shown in Figure 3. Several novel applications are discussed below. Microchannel contractors allowing the intensive contacting of two immiscible fluids and thus facilitating separations, e.g., by extraction, were recently developed [33]. Typical channel dimensions were in the order of 100 µm for channel heights; ranging 10 to 50 µm for thickness of the contactor plate; and between 1 and 20 cm for channel length. Note that while the application of the contactor plate facilitates countercurrent operation, it also increases mass transfer resistance. This

Figure 3 Example of an integrated microreaction system consisting of heat exchangers and mixing unit operations. (From Ref. 67.)

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resistance can be decreased by reducing the thickness of the plate, increasing its porosity, or reducing its tortuosity. This last can be attained by drilling cylindrical or conical holes in the thin substrate materials. Mass transfer performance is basically characterized by three parameters. The first performance parameter is the inverse Peclet number for mass transfer. The second performance parameter is the residence time. The third parameter, termed as the breakthrough pressure, expresses that the maximum allowable pressure drop across the contactor plate is limited. Experimental results showed that the size could be reduced by as much as 2 orders of magnitude compared to usual sieve tray columns. The control of fluid flow in microfabricated devices raises several difficult problems, especially in the case of disposable or mobile biochemical motherboard systems. An innovative design for controlling fluid flow is based on the application of a structurally modifiable fluidic network of passive microvalves (restrictions), which allows the flow sequence primarily controlled by the microvalves. Modifying the relative values of the passive valves with respect to channel geometry or their arrangement enables programming the fluid delivery sequence in the microfabricated network [34]. The utilization of microfabricated systems to cultivate, transport, and manipulate living cells has been reported by several groups [35,36]. A microfluidic device made of polydimethylsiloxane (PDMS) applied for the growth of bacterial cultures successfully demonstrated the feasibility of monitoring and cultivating the microbes in such systems. Potential benefits compared to conventional devices include the possibility of real-time monitoring, higher sampling frequency, simpler automation, and reduction of specimen volume and biological waste [37]. 14.3 ANALYTICAL ASPECTS AND APPLICATIONS Microfabricated devices that require new approaches to sensor and detection technology in order to utilize the full potential of microsystem integration, on the other hand, provide complete solution to complex and tedious analytical problems.

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14.3.1

Chemical Sensors

Successful implications of microreaction technology should include appropriate detection techniques, e.g., chemical and biochemical sensors, fluorescent reporter techniques, spectroscopic methods, mass spectrometry (MS), nuclear magnetic resonance (NMR) spectroscopy, etc. Quantitative and qualitative analysis of chemical and biochemical products is usually a labor-intensive and time-consuming part of the research and development process. Novel approaches such as photoacoustic spectroscopy (PAS) provide a simple and general optical method for gas-phase detection. In photoacoustic (also called optoacoustic) spectroscopy, incident light is modulated at an acoustic frequency. If the optical wavelength couples to any of the energy transitions in the gas, the gas absorbs the light, resulting in a periodic gas expansion [38]. The potential and feasibility of photoacoustic spectroscopy for general microscale chemical analysis are illustrated by the detection of C 3H 8 in a photoacoustic cell that is implemented in a microfabricated chemical reactor and also by microfabrication of a photoacoustic detection cell [39]. Jensen’s group [40] incorporated online detection into a micromixer-based liquid-phase reactor allowing the application of visible and UV spectroscopy. SU-8 (an epoxy-based, negative resist), composite quartz/SU-8, and silicon materials were used for constructing the microfluidic channels. In addition to realizing chemical functions, microfabrication procedures can also be used to integrate micro-optical components into the microchip. Such microoptical elements readily provide necessary optical functionalities such as illumination, collection, and signal filtering, all required for highsensitivity fluorescence detection comparable to those attained with standard detection systems. Dandliker and coworkers [41] reported a completely integrated microfluidic/micro-optical device, with refractive microlens arrays and aluminum aperture arrays deposited on both sides of a microfluidic chip. 14.3.2

Microanalytical Devices

In the early 1990s in Basel, researchers in the Widmer lab proved the usefulness of electrophoresis microchips [42]. Other groups followed the lead and applied microfabricated device technology to the analysis

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of amino acids [43], DNA restriction fragments [44–46], PCR products [47], and DNA sequencing [48]. Figure 4 depicts a microchannel design applied for rapid PCR, preconcentration, and analysis [49]. Similar to microreactor technology, electrophoresis microchip-based separation devices were also developed using techniques elaborated by the semiconductor industry, suggesting that channels and other functional elements can be fabricated in glass substrates by microlithography. With the advent of the so-called double-T injector setup, well-defined amounts of samples could be readily analyzed on electrophoresis microchips [50]. Samples are typically loaded electrokinetically into the injector region, then the analyte molecules are separated by means of applying the electric field not only across the separation channel but

Figure 4 Microchannel structure design for integrated rapid PCR analysis. On-chip sample preconcentration is achieved by means of an incorporated porous membrane structure in the injection valve. (From Ref. 49.)

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Figure 5 Design layout and scanning electron micrograph image of a micromachined chromatographic phase support. (From Ref. 53.)

in a smaller extent to the sample and waste reservoirs as well, to prevent bleeding of the sample into the separation channel [51]. The separated solute molecules are then most frequently visualized by confocal microscopy with laser-induced fluorescence (LIF) detection. Short injection plugs, high electric-field strengths, and short effective separation lengths result in rapid separations in seconds and extremely high separation efficiencies because of the minimized extra-column broadening effects [52]. Microfabrication also offers a novel approach to highperformance micro- and nanovolume liquid chromatography through the application of micromachined chromatographic phase support [53]. Figure 5 illustrates the layout and the scanning electron micrograph image of such a micromachined chromatographic support. 14.4 MICROFABRICATION TECHNIQUES Based on the fact that silicon microchips provided an ideal subject to miniaturization in the electronics industry, including the possibility of even increasing capacity and functionality, microfabricated chemical and biochemical reactor devices are manufactured from a variety of materials applying similar processes. Besides the commonly used glass and silica wafers, polymer-based materials such as polydimethylsilox-

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ene (PDMS), polymethylmethacrylate (PMMA), polycarbonate (PC), and Teflon have been recently introduced, all of which are especially suitable for screening catalytic reactions and drugs and also for a wide range of applications. Some of the most important fabrication methods are microlithography (soft and photolithography), molding, embossing, and micromachining applying laser, electrochemical, or ultrasonic technologies [54]. Another microfabrication technology, called LIGA, combining lithography, electroplating, and molding, was successfully applied to the production of microreactors and other microdevices. This technology allows the application of any conductive surface materials and enables injection molding [55]. Metal devices are often employed in case of fast exothermic, heterogeneously catalyzed reactions and also in different separation processes at pilot and production scales. Metal devices are generally fabricated by microlamination techniques with thin laminated metallic layers forming channels and partitions. Pressure stamping or photochemical machining techniques are used to provide the necessary patterning [9]. 14.5 SIMULATION OF MICROREACTOR SYSTEMS Computational fluid dynamics (CFD) simulation can provide valuable insight into a plethora of liquid phase processes, such as mixing, fluid transport, separation, etc. [4]. Computer simulation is primarily considered as a design tool but is also used to support the interpretation of experimental data. A general, steady-state, finite element approach to simulate two- and three-dimensional fluid flows, thermal fields, and chemical concentrations in microfabricated devices was based on the Galerkin finite element method [56]. The model system consisted of two coupled domains, the flow channel and the surrounding solid material exchanging heat with the flow channel. This simulation method facilitated the exploration of various microdevice designs also demonstrating the abilities of computer simulation during the development of membrane-mediated systems [57]. The macroscopic description of electrokinetic transport in microfabricated channel networks is based on the principles of multicomponent continuous media mechanics, coupled with equations describing the electric field [58]. The frequently quoted ‘‘Lab-on-a-Chip’’ approach can address ever more complicated

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chemical and biochemical problems, and will necessitate correspondingly sophisticated fluidic design and complex electrokinetic control methods. In addition to solely relying on experimentation to develop microfluidic structures, computer-aided design and analysis tools are more and more explored for prototyping and refining fluidic arrangements. A significant reduction is expected in both time and expense when such tools are implemented. The principal transport mechanisms of electric field–mediated migration (electrophoresis and electro-osmosis) and diffusion should be considered during the design of microfabricated devices. A comprehensive computer program, simulating the mass transport in any twodimensional channel structure was developed by Ermakov et al. [59]. Their first approximations considered uniform electric conductivity, insignificant Joule heating, and constant thermophysical and surface properties. These assumptions resulted in a partially independent flow characteristics, electric field, and concentration distribution. A rectangular grid-based finite-difference algorithm was employed to solve this simplified model. Computer simulation assisted the examination of different aspects of operations and helped to reveal optimal operation parameters for various cross-channel-based injection methods (gated and pinched type) [60]. Computer simulation of pinched injection in a cross channel structure is illustrated in Figure 6 [61]. It was also demonstrated that such basic microdevice elements as the simple T-structure and channel-cross readily accommodate sample mixing and electrokinetic focusing, respectively [62]. A threefold approach was applied for modeling a microfluidic separation chamber for the isolation of various cell types from a heterogeneous population, based on their dielectric properties. Fluidic characteristics were optimized applying the computational fluid dynamics package STAR-CD. The Ansoft Maxwell 2D field simulator was employed to simulate the various electric field–mediated forces within the device, enabling electrode geometry optimization. Effectivity of cell separation from various mixtures was modeled prior to microfabrication, using the MATLAB software package [63]. The accurate performance simulation and detailing of microfabricated separation devices offered by simulations can provide efficient system design and optimization schemes. Accurate flow and thermal predictions are also enabled

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Figure 6 Simulated images and the electric-field distribution during pinched injection in a simple cross structure. (From Ref. 61.)

by the lattice-Boltzman simulation method for microfabricated devices even in complex geometrical setups. Additional terms such as surface forces can also be included to simulate discrete liquid-vapor and liquidliquid phase interfaces, as well as surface adsorption properties [64]. Focusing on new applications and requirements of the biochip industry, analyses of electrokinetic injection phenomena [65] and prediction of electrophoretic motion in microfabricated channels [66] were recently reported, using a commercial finite element microdevice simu-

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lator, FlumeCAD (Coventor, Inc.). This highly sophisticated software package enables general design of integrated chemical and biochemical processes and/or analysis systems, also supporting three-dimensional simulation of chemical transport in electrophoretic, electro-osmotic, and pressure-mediated systems.

14.6 DESIGN AND PARALLELIZATION OF MICROFABRICATED REACTORS The significantly improved mass transfer characteristics in micro-unit operations compared to corresponding macro-scale devices can mainly be attributed to the large gas-liquid interfacial area maintained in these systems [4]. One of the most critical aspects of microreactor design is to counterbalance the possible increase in pressure drop caused by the reduced dimensions by the gain in heat and mass transfer. Dividing the flow into a number of channels increases the effective cross section area, resulting in reduced pressure drop, providing high reactor throughput, but still maintaining the high surface-to-volume ratio. In case of reactions limited by mass transfer processes, the volume of the microreactors can be significantly smaller while high productivity is still retained. Using scale-out methodology to provide the required processing capability would help to eliminate problems arising from conventional scale-up design approach. The scale-out philosophy together with an appropriate large-scale microfabrication technology enables one to extend reaction conditions, optimized on a single device, by applying an adequate number of single units referred to as ‘‘parallel scale-out’’ [9].

14.7 CONCLUSION Chemical and biochemical microreactors represent a novel approach in respect to production flexibility also applicable to large-scale production by parallelization. Some of the main advantages of miniaturization are reduced reagent consumption; improved-performance, interconnected network of channels with multifunctional possibilities; inherent mechanical stability of monolithic systems; possibility of paralleliza-

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tion; and inexpensive mass production. Miniaturization greatly improves heat and mass transfer; it opens new horizons and revolutionary changes in chemical processing and biotechnology. System integration also plays one of the most significant roles in miniaturization. As the new millennium holds great promise for the development of microfabricated chemical and biochemical systems, the quest for miniaturization is going to lead to greater process intensification.

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15 Approaches to Miniaturized High-Throughput Screening of Chemical Libraries Damien Dunnington, Zhuyin Li, Alan Binnie, and Henning Vollert Aventis, Bridgewater, New Jersey

15.1 INTRODUCTION Over the course of the past 30 years, the process of searching for new drugs has undergone both explosive growth and fundamental change. One of the consequences of this shift is the requirement for assay miniaturization. The vast potential profit of marketing novel drugs has recently driven the pharmaceutical industry to accelerate and intensify the initial stages of discovering new drugs (known collectively as ‘‘lead generation’’). Discovery of a new drug via high-throughput screening (HTS) begins by associating chemical compounds with a desirable biological activity (a ‘‘target’’). In an effort to maximize opportunities for success, the industry has increased both the number of targets screened per year and the number of compounds tested in each screen. Thus, rather than testing a few hundred compounds as potential drugs in a few animal disease models per year, drug discovery research at the typical large pharmaceutical company currently entails testing hundreds of thousands of compounds for desirable activities in dozens of 371

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different in vitro biological assays each year. This trend is certain to continue, as both the number of potential targets and the number of compounds available for testing are multiplying quickly. Advances in the human genome project, proteomics, and bioinformatics are providing new targets at an unprecedented rate; in fact the number of potentially attractive targets far exceeds the capacity of the industry to pursue them. At the same time, recently developed combinatorial chemistry techniques are causing an exponential increase in the number of compounds available for testing. In order to meet this dramatic increase in demand for screening, the concept of ‘‘ultrahigh-throughput screening’’ (uHTS), defined as screening ⬎100,000 data points per day, has become increasingly popular [1–4]. The increased cost associated with screening more targets and more compounds has pushed the industry to search for ways to decrease cost, and one of the most direct and effective means to achieve cost savings is to perform uHTS in much smaller volumes. Although it is technically possible to attain uHTS in 96-well plates, the costs associated with massive screening in 96 well plates are prohibitive. For this reason, the concept of assay miniaturization is closely tied to that of uHTS. The term ‘‘miniaturized assay’’ is commonly defined as an assay performed either in microtiter plates of 1536-well or higher density, or in volumes of ⬍5–10 µL. The trend to reduce assay volumes is not new, and has been apparent ever since the advent of pharmaceutical screening. The same forces that have driven the continuous reduction in assay volumes over the decades are now driving the movement into miniaturized assay formats. The most obvious incentive to miniaturize comes from reagent savings, since many of the components of screening assays are extremely expensive. Other benefits include improved throughput in detection, reduction in waste flow and storage space, and the reduced need for labor and automated screening systems. In vitro pharmaceutical screening was originally conducted in test tubes, with the assay material being transferred to cuvette-based instruments for detection. Although 96well microtiter plates have been available since 1960’s, they were not initially used for pharmaceutical screening. The initial application of microtiter plates was restricted to enzyme-linked immunoabsorbance assays (ELISA), and detection was most commonly achieved by a qualitative visual assessment of colorimetric readouts. True HTS as we

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know it today began in the 1980s, when equipment capable of optically quantifying signals directly in 96-well plates and multichannel pipetters became commonly available. In response to the demand for microtiter plates with greater sample density and lower effective working volumes, manufacturers introduced the 384-well plate in early 1990s. The industry has been able to adapt to the 384-well standard fairly quickly because the liquid handling systems and the assay technologies in use for 96-well screening are, in most cases, directly applicable to the new format. The rate limiting factor in the 96- to 384-well conversion has been the need to acquire updated detection systems, as most of the original 96-well plate readers were not compatible with 384-well plates. Almost all modern plate readers are now compatible with the 384-well format. 15.2 COST SAVINGS AND THROUGHPUT Although there is universal agreement throughout the industry that assay miniaturization will lead to significant cost savings, estimates of the magnitude of savings to be derived from each component in the overall cost varies widely. Cost components include compound samples, biological reagents, consumables (plates and tips), waste disposal, equipment costs, and labor. 15.2.1

Compounds

HTS compound samples fall into several categories, and the costs associated with acquisition, registration, storage, and retrieval for each type of sample are different. The price of acquiring samples varies widely, depending upon the type of sample. Historical compound collections are built over long periods of time as a byproduct of the pharmaceutical business, and are recycled back into new projects for screening. Combinatorial chemical samples are produced either internally or acquired externally. In the case of internal development of synthetic methods for strategic compound families, the development costs, as well as the cost of the building blocks used, may be extremely high. Natural products samples include extracts from plants, fungi, bacteria, and marine organisms. Chemical HTS samples acquired on the open market range

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in price from $5 to $100/mg, averaging about $10/mg when purchased in bulk. Using $10/mg as a benchmark, for a screen of 1 million compounds at 10 µM in 100 µL in 96-well plates and assuming a 10-fold waste factor due to intermediate dilution, compound acquisition contributes only $50,000 to the estimated average of $800,000 total cost for the screen. Using this calculation the impact of miniaturization on compound costs in minimal, because the compound is less than 10% of the total cost. At the other extreme, an estimate of $250,000 to $1 million has been quoted [2] for the same amount of screening using combinatorial chemistry samples and figuring in chemical development costs. Given that most of the cost resides in the synthesis procedure, conservation of library compounds through miniaturization can lead to long term savings by increasing the number of targets that can be screened per sample lot. Additionally, combinatorial chemistry and automated synthetic procedures produce limited amounts of material, and reduction in usage of such samples is clearly beneficial. 15.2.2

Bioreagents

In the average assay, the greatest saving from assay miniaturization will be realized in bioreagent costs. Included in this category are incidentals such as scintillant and reagents used to coat assay plates. Estimates for the cost of bioreagents vary tremendously. Industry estimates for 100 µL 96-well assays range from a low of about 2 cents per well for readily available enzymes up to $8 per well for new technologies such as SPA beads. Reducing the volume of a million compound screen from 100 µL in a 96-well plate to 2 µL in a 1536 plate thus saves between $19,600 and $8 million per target, depending on the type of targets. Of course a 96-well assay costing $8 million in reagents would not be performed today, but reducing the cost to $160,000 via miniaturization would bring the cost into the realm of feasibility. 15.2.3

Consumables

The cost of pipette tips and the plates themselves are a relatively minor component in most assays. However, once the other costs are scaled down, the cost of the assay plate itself becomes a significant portion of the total cost. Standard 96-well plates currently cost between 50

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cents and $2, and 384-well plates average about $4. At this point in time, due to the limited market and attempts to recuperate development costs, 1536-well plates cost as much as $30 each. This situation will undoubtedly improve when more manufacturers enter the market, as competition will force the price closer to the actual cost of manufacturing the plates. 15.2.4

Waste Disposal

With the exception of some radioactive assays, the cost of waste disposal is negligible. Most assay materials are considered non-hazardous and are discarded in the normal waste stream (even radioactive assays, in Arizona). However, this cost may be quite high if the screening site is located in an area with strong regulations. The drive to miniaturize has in turn led to increased usage of fluorescence technologies and more widespread acceptance of alternatives to radioactivity. 15.2.5

Equipment Costs

The impact of miniaturization on equipment costs is frequently underestimated. Because all of the equipment is durable (but may become obsolete and may be based on unproven technologies and need frequent upgrades and/or repairs.), these capital expenditures are only incurred once and thus are averaged over the price of many screens. The main areas to be considered are the cost to equip assay miniaturization labs, the cost of purchase and maintain robotics screening systems, and the cost to store and handle compound samples. Although at present the initial cost of screening equipment for miniaturized assays is higher than that for 96- and 384-well use, the price gap is already beginning to close. Because of the higher density of miniaturized assays, the number of robotic systems required to screen will be reduced. An organization that wishes to screen 100 targets per year with 1 million compounds requires a capacity of 100 million wells per year. Using a linear track robotic system (with a total capacity of about 180 plates per day) as a benchmark and assuming 150 24-hour working days per year, the organization would require 39 systems, each costing close to $500,000 to complete this task. Assuming a direct scaling of tasks, the same amount of screening could be performed on three 1536-enables sys-

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tems. Although the initial setup costs of the 1536 system would be slightly higher, the increase would be inconsequential compared with the reduction in cost due to less system redundancy. Finally in the long term, storage and retrieval of compound collections in 1536 plates would attain large savings in system costs, but this would require concomitant reductions in the amount of compound needed for primary and secondary assays. Compound storage and retrieval systems capable of holding and tracking 1 million compounds currently cost between $2 million and $15 million. Reducing the number of plates by 16-fold will undoubtedly result in large savings in this area. 15.2.6

Labor

The net effect of assay miniaturization on labor costs is difficult to predict at this time. Although fewer personnel will be required for screening, these savings may very well be offset by the need for more effort in assay development. Because many of the most convenient miniaturized assay types are more difficult to develop than traditional types, assay development challenges may be a recurring facet of overhead associated with assay miniaturization. 15.2.7

Beginning the 1536 Transition

The transition from 384- to 1536-well format is just now becoming feasible. The first commercially available 1536 plate came to market in 1997, followed in 1999 by the commercialization of a low profile 1536-well plate [5]. The first 1536-compatible plate readers and parallel liquid handling devices appeared on the market in 1998. Yet with all of the components for 1536 screening in place for almost two years, only a few laboratories have managed to implement true HTS screening in the new format. Although proof of concept for miniaturized formats of most common assay types is straightforward when performed manually in a few wells, progression to screening of full compound collections while maintaining sufficient quality is still a considerable challenge. A survey of HTS directors [6] reveals that despite the remaining obstacles, 1536-based screening is planned for about 10% of the total industry screening effort in 2000.

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15.3 NEW TOOLS FOR MINIATURIZED SCREENING SYSTEM The trend in assay miniaturization has created an increasing demand for microscale liquid handling, ultra sensitive detection and versatile automation systems, as well as appropriately engineered assay plates. As a result, many technologies have emerged to address these unmet needs for HTS. 15.3.1

Pipetting Systems

This section reviews the new liquid handling technologies that are able to dispense or to transfer nanoliter or even picoliter volumes. It should be noted that due to the recent emergence of these technologies, and that this field is developing primarily outside the academic world, this article contains little published literatures and is based on product literatures, conference proceedings, customer applications and personal communications. With respect to miniaturized pipetting and dispensing systems there are three main technologies on the market: (1) piezoelectric and inkjet systems; (2) air displacement systems; and (3) pintool systems. Piezoelectric and Inkjet Microdispensing Systems Piezoelectric and inkjet systems were the first systems on the market that were able to dispense submicroliter volumes [7]. Inkjet microdispensing is derived from the inkjet printing industry. The principle of inkjet dispensers involves the application of pressure created by a syringe to liquids. Liquid droplets then pass through a small orifice and eject into wells upon valve opening. In general the volume of the droplets is in the range of 1 nL to a few hundred nanoliters. Furthermore, droplets can be dispensed into small wells without the tip entering into the well. Unlike inkjet dispensers, piezoelectric dispensers utilize a piezoelectric crystal which is in contact with a micromachined capillary or silicon/glass cavity. Applying a voltage across the piezo causes the wall of the capillary or silicon membrane of a cavity to deform, and subsequently eject droplets. In general, piezoelectric dispensing sys-

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tems can dispense drops between picoliters and a few nanoliters. The systems operate at rates up to 100 to 2000 drops per second. Because the ejection speed of the droplets is in the range of several meters per second, mixing does not present a problem for 1536-well plate formatted assays provided aqueous solutions are used. Despite their high accuracy, piezoelectric and inject microdispensing technologies have not been widely accepted in HTS. The system is prone to clogging by fine particles and gas bubbles (especially piezoelectric systems). Secondly, the actual volume dispensed depends on surface tension parameters of the reagents. Thus, calibration is necessary for each new reagent. Thirdly, a carrier fluid must be used during aspiration and dispension which results in high reagent waste. Finally, the compatibility of these systems for dispensing vesicles or cells, especially mammalian cells, needs improvement. Air Displacement Systems Apart from piezoelectric and inkjet technologies that eject nanodroplets, several companies have miniaturized their standard air displacement pipetting systems (Fig. 1A). These systems are based on syringe pumps or pistons with working volumes between 0.5 and 20 µL (utilizing bigger tips: between 4 and 250 µL). Compared with microdispensing systems, the air displacement instruments have a significantly higher working volume. Provided a final assay volume between 2 and 10 µL is satisfactory, these simple machines may be an efficient choice. Several instruments on the market have 96 or 384 pipetting channels. They take less than a few minutes to transfer compounds from four 384 plates into one 1536 plate. In addition, these instruments are familiar and easy to use. The fact that these instruments and software packages are similar to standard pipetting systems should not be ignored, as the throughput of a screening lab closely depends on fast troubleshooting. Although several companies claim to have miniaturized air displacement systems in their portfolio, only a few have actually delivered robust instruments that offer precision of ⬍10% cv. This precision is highly dependent on the exact alignment of the pipetting head and the plate feeding system. Furthermore, the performance of the machines crucially depends on the quality of the tips and plates. Curved plates

(A)

(B)

Figure 1 Liquid transfering tools: (A) 384-channel air displacement pipetting systems; (B) 384-channel pin tool.

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(the difference in height between the corner and center of the plate should be ⬍250 µm) can cause significant pipetting errors which may lead to a high number of false-positive values. Nevertheless, the first big pilot screens and the first extensive routine screens were performed utilizing these miniaturized low-tech machines. The future will determine if piezoelectric/inkjet systems or miniaturized air displacement systems conquer the screening laboratory. Pin Transfer Technologies Pin transfer technologies rely on the use of metal pins (Fig. 2B) to transfer liquid from a reservoir into wells of a microplate [8]. The pin is dipped into a liquid source plate (e.g., chemical compound or reagent), and a small drop sticks on the tip of the pin due to surface tension. Subsequently, the pin is moved to the microplate and touched to dry surfaces or into prefilled wells. Typically, pin transfers are between 100 nL and a few nanoliters. Although the precision of the pins is limited (⬃10–30% cv), the transferred volume is fixed and dependent on the surface properties of those liquids transferred, pins have the advantage of being inexpensive, robust, and can be arranged in 384- or 1536- channel arrays. Unlike piezoelectric or inkjet systems, the pin does not require dead volume: small volumes can be transferred directly out of storage plates without any dilution steps. This significantly reduces the consumption of chemical compounds and other consumables, and enables the direct transfer

Figure 2 A telecentric lens is less prone to lateral effects that include parallax error and loss of photons than the standard lens.

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of compound from master plates to assay plates, thus increasing the throughput of screens. Therefore, utilization of pins for compound transfer is a possible alternative if cost and throughput are more important than precision and volume accuracy. 15.3.2

Detection Systems

Assay miniaturization calls for highly sensitive detection systems capable of analyzing results in 1536-well plate formats and beyond. Miniaturized standard plate readers, imaging systems, and single molecule detection systems are the three major available technologies that meet these challenges. Miniaturized Standard Readers As with the miniaturized standard pipetting systems, several companies miniaturized their standard photomultiplier tube (PMT) based plate readers. These detection systems analyze one well after the other. This of course is a time-consuming process but fortunately there are instruments on the market that take only 1–8 min to analyze the prompt fluorescence of a 1536-well plate. Some of the systems can adjust the z-height automatically, and thus obtaining optimized signal to background ratio, and having detection limits lower than 1 nM for fluorescein. These systems are readily integrated into the HTS laboratories because the companies can use their standard plate feeding system, and they can be operated easily by technical staffs. Digital Imaging Digital imaging, which utilizes a charge-coupled device (CCD)-based detection technology, is another type of detection system now available for assay miniaturization [9]. The development of high-performance CCD cameras and telecentric lenses (Fig. 2) has made it possible to analyze assays performed in various high-density microplate formats [2,10]. The attraction of imaging systems lies in the ability to read miniaturized formats of virtually any type and the speed of detection. An area imaging system detects the entire microplate in one step and therefore the analysis can be very fast. Because of the high speed, im-

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ages of the whole plate can be taken in a rapid sequence. This could be the basis for the analysis of dynamic assays. There are greater technical obstacles to imaging systems than to plate readers. In order to achieve sufficient sensitivity, CCD cameras are cooled down to ⫺45°C (or lower). At low temperatures, the noise of the CCD chips is reduced significantly. Besides cooling the system, a dark noise correction needs to be performed to subtract the electronic noise of the camera and to increase the dynamic range of the camera. The dark noise correction depends on the exposure time, and correction is recommended at an exposure time close to the sample exposure time. Furthermore, intensive standardization of the imaging system is required. Although telecentric lenses are used, all commercially available CCD cameras view lateral wells at an angle, therefore reducing the brightness of these wells. The brightness of wells in the center of a plate is higher than that of the wells at the corners, making a flat field correction necessary to standardize the results. To date, an HTS imaging system which utilizes automated plate handling and is compatible with 1536-well plate is not available in the market, therefore the throughput of plates reading is limited for the imaging system. Another limiting factor is that the systems can analyze only a few readout methods in 1536-well plates (fluorescence, luminescence, absorbance, certain isotopes). In spite of these and other technical hurdles, several feasibility studies and pilot screens have shown that imaging systems generate similar results, identify the same active compounds, and yield similar IC 50 data compared to plate readers [11– 13]. The performance of luminescent assays in particular has shown that cooled CCD cameras permit fast screening of miniaturized cellbased assays. Compared to standard luminescence plate readers, cooled CCD cameras are about five to eight times faster [14]. Scanning Confocal Imagers An alternative to bulk field imaging is to scan a finely focused laser beam across the well or plate surface and measure reflected light or fluorescence emission. A high resolution image can be constructed by ‘‘rastering’’ the beam across the image field. Instruments that accomplish this are in early stages of development for HTS applications and are well suited to miniaturized assay formats. In particular, such ap-

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proaches can be used to measure compartmentalization of fluorophores within cells, providing access to mechanistic information on the effects of compounds. However, there are a number of practical issues that must be solved before scanning imagers can become sufficiently robust for routine HTS usage. The base of most microtiter plates is not perfectly flat and a tracking mechanism is needed to align the optics with the plane of the assay reaction, especially when cells are being imaged. Software development is needed to extract meaningful quantitative information from the mass of data produced during imaging. Read times must be compatible with the duration of reactions or cellular translocation events, and high speed scanning systems may be needed to capture transient phenomena. Despite these needs, the sensitivity and resolution of scanning imagers provide a means to extend the range of assays that can be accommodated in miniaturized HTS formats. Single-Molecule Detection Systems Based on Confocal Optics The ultimate goal of high-sensitivity detection schemes is observation at the level of a single molecule [15]. The detection of single molecules is achieved by utilizing confocal fluorescence methodologies. These techniques are based on detection of laser-induced fluorescence of dyetagged single molecules in a very small focal volume element of ⬃1 fL. Among confocal methodologies, fluorescence correlation spectroscopy (FCS) is a well-established method. Several research groups in the academic world have used this technology for ⬃20 years [16,17], and for the last few years it has also been used in the high-throughput screening [18–20]. Fluorescence correlation spectroscopy is a technique that allows the determination of the diffusion constant of a fluorescent molecule in solution. Also, the binding of the fluorescent molecule to a target can be analyzed if the difference in the diffusion coefficients of the free and bound ligand is sufficiently large. This technology is very sensitive (it detects single fluorescent molecules); however, the analysis speed of FCS is limited—especially when utilizing large-target molecules or cells. The limited throughput is still the main bottleneck of the FCS screening approach. In the first pilot screens utilizing a simple homogeneous assay the FCS took ⬃1–2 sec/well. Thus, at least the

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screening of non-cell-based assays can be achieved at the highthroughput screening level. To a certain extent, the low throughput will be compensated by the development of multichannel reader systems. A further development of FCS is dual-color fluorescence crosscorrelation spectroscopy (dual color FCS). In dual-color FCS the concentration and characteristics of two fluorescent species, as well as their reaction products, can be followed in parallel. By using two differently labeled reaction partners (e.g., a ligand labeled with one dye and a receptor labeled with another dye) the selectivity to investigate the temporal evolution of reaction product is significantly increased compared to one-color FCS. It has been shown recently that the dual-color FCS improves FCS detection methods in terms of analysis speed, specificity, and sensitivity [21,22]. 15.3.3

Plate Formats

Because assays have proceeded into smaller and smaller volumes, one must ask whether the 1536-well format, with its current minimum assay volume of ⬃2 µL and maximum of ⬃10 µL, offers sufficient cost saving, or whether higher densities are desirable. Plates with densities ranging from 3456 [23], to 6144 and 9600 wells [10] and CD-type plates [24] have been produced and used for assay development. For a variety of reasons, it seems that 1536 may remain the industry standard, at least for the next few years. For the sake of simplicity, as well as to reduce engineering problems for the manufacturers of liquid handling and detection equipment, it is desirable to have a single standard plate type for all assay types. The plates should be compatible with existing screening equipment, meaning that the overall footprint should stay the same as 96- and 384-well standards. The plates should be compatible with existing 384-channel pipetters, which eliminates all of the formats except 1536 and 6144. In order to accommodate the volume required to culture sufficient quantities of cells for assays and the additional volume required to lyse cells prior to detecting intracellular reporter enzymes, the lowest desirable well capacity is ⬃3–4 µL. Absorbance assays with a low concentration of chromophore also demand a longer path length for sensitive detection. The 6144 format is too closely spaced to accommodate these volumes, and thus is not a

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generally applicable plate type. Perhaps the most compelling argument for remaining in the 1536 format is that of opportunity costs. The equipment to perform assays in the 1536-well format, although not perfect, will enable cost effective uHTS in the very near future. The assay types common across the industry, almost without exception, can be implemented in 1536 without extensive redevelopment. Moving to assay volumes ⬍1 µL would require significant new development in both assay techniques and equipment, causing a significant delay in realizing the increased throughput and cost savings of assay miniaturization. 15.3.4

Automation

According to plate transport method, automation in HTS can be classified into two concepts: workstation and fully integrated robotic system [25]. Both methods can be applied to conduct screening in miniaturized formats. Using the workstation method, plates are transported manually between different functional stations which are sometime integrated to stacker or feeder unit. It has the advantage of being very high throughput for individual assay steps, allows multitasking several stations, is economical, provides flexibility, and requires less laboratory space. However, workstation methods require continuous human supervision and provide only linear access to individual stations. In addition, it is inconvenient for complex assays and for kinetic readouts. Fully integrated systems comprise of pipetting stations and other functional modules integrated with one or several robotic arms. These systems have sophisticated scheduling software programs that allow for unattended runs with nonlinear access to different functional units. Multistep assays or kinetic readouts can easily be programmed at the robotic system; different functional units by different manufacturers can also be integrated into one system. Compared with workstations, some integrated robotic systems have slower plate handling and have troubles due to robotic arm crashes; the plate movement of the arm can sometimes be the rate-limiting factor in the throughput of the assay screen. In addition, the systems are costly, they require large laboratory space, specialized expertise and service support. Sometimes, the functionality that is required in an assay may not be available on the integrated system. In general, the decision for which approach to take is dependent

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upon the preferred methodology of the screening department, the number of the assays, the assay complexity, the throughput desired, the space and budget available. 15.4 ASSAY DEVELOPMENT FOR MINIATURIZED SCREENING The increasing demands for miniaturization and automation in the HTS environment call for robust, reproducible, scalable, and standardized assays. Furthermore, these miniaturized assay formats should still generate high quality data that reflect the true activity of the biological targets. Consequently, the miniaturized assay method of choice is dependent on the type of target, the desired outcome of the screen, the properties of the compound library, and instrument and reagent availability. To expedite assay development, early communication between the assay originator, developer, and screener is critical. Once the target is selected and validated, the team should define the fundamentals of the assay. This includes whether the assay will be biochemical or cell based, if the outcome is an agonist or antagonist, and whether the antagonist is a competitive or non-competitive inhibitor. Knowledge about the target at the molecular level in terms of its sequence, structure, signal transduction pathway, thermodynamic and kinetic constants, the chemical library in terms of combinatorial or discrete, color intensity, solubility, and cell permeability, and reagents in terms of availability, protein purity, and cell line type, will be necessary to design an optimal assay method. Homogeneous assay formats, where reagents are mixed and read without separation or washing steps, is the preferred choice for miniaturization purposes. The development of miniaturized heterogeneous assays where mixing, separation and washing steps are required, remains a challenge. Advancements in micromechanical and microfluidic technologies will provide solutions for such problems in the near future. Currently, many pharmaceutically interesting biomolecules, such as receptors and enzymes, are screened using a combination of radioactive ligands and scintillation proximity technologies in pseudo-homogeneous formats [26]. However, in the absence of new and improved detection methods, miniaturized assays based on radioactivity are im-

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practical. For example, to scale down a conventional kinase assay which measures the ability of the enzyme to transfer (gamma-33 P)phosphate from 50 µL to 5 µL, the counting time per well would increase from 10 sec to 10 ⫻ (50/5)2 sec to achieve the same precision. Although parallel detection methods, such as imaging systems with CCD cameras, have been developed to speed up the counting process, problems with tracer-specific activity, detector sensitivity, health and environmental safety, and screening costs have prevented assay developers from further miniaturizing radioactive-based methods. The desire to get away from radioactivity has lead to the development of new assay technologies where signal detection is based on optical methods such as fluorescence, luminescence, and absorbance [10–22,27–33]. Among these, fluorescence techniques are the most popular. At a given sensitivity requirement, fluorescence based detection methods are more easily miniaturized without sacrificing precision when compared with radioactive and other optical methods. First, each fluorophore can be excited many times during the illumination period before it is photo bleached. Second, average fluorescence intensity is dependent on excitation energy, therefore a powerful laser illumination technique can be used to increase the population of excited fluorophores. Unlike luminescence techniques in which weak signals start to decay immediately following reagent mixing, fluorescence signals are relatively stable during the detection period. For these reasons, many fluorescence based detection techniques have been developed to accommodate different needs within miniaturized high throughput screening. 15.4.1

Biochemical Assays

Most biochemical assays where targets are enzymes, nuclear or membrane-bound receptors, or protein-protein interaction pairs can be configured into miniaturized fluorescence-based homogeneous assays that mimic physiological conditions. These assay formats are usually based on proximity or mass-dependent principles to measure molecular interaction. Examples of proximity-dependent assay methods are fluorescence resonance energy transfer (FRET), its elaboration homogeneous timeresolved FRET (TR-FRET/HTRF) and lifetime-discriminated FRET

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[34–39]. Proximity-dependent principles can be applied to the study of proteases, receptors, protein-protein interactions, and protein-DNA interactions. For example, proteases, which catalyze the cleavage of specific peptide bonds, can be studied using FRET methods (Fig. 3). A donor dye is attached to one side of the cleavage site, and an acceptor, which may be a fluorescent or a nonfluorescent quencher, is attached to the other side. The distance between dyes must be close enough to allow resonance energy transfer to occur without interfering with enzyme activity [40,41]. Reactions are monitored at the donor’s emission wavelength. The proteolytic cleavage of the substrate separates the donor from the acceptor, thus disrupting the energy transfer and resulting in an increase in fluorescence intensity. Mass-dependent assay methods include fluorescence polarization (anisotropy, FP/AP), fluorescence correlation spectroscopy (FCS), and their time-resolved or lifetime-discriminated variations. Mass-dependent effects can generally be utilized to study proteases, transferases, and receptors [18,30,31,42,43]. For example, a polymerase, which transfers subunits onto macromolecules, can be assayed by the FP based method. By labeling the subunit with a fluorescent tracer, the

Figure 3 FRET-based protease assay.

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transfer of the subunit to the macromolecule can be monitored through a fluorescence polarization change. In the absence of molecular transfer, the fluorescent tracer rotates quickly and the resulting emission is depolarized. Upon the transfer of the subunit to the large macromolecule the tracer remains relatively stationary during the lifetime of the fluorophore, and the emitted light will have a high degree of polarization. The time-resolved and lifetime-discriminated fluorescence techniques can greatly reduce assay variation caused by light scattering from membrane preparations, compounds or labware, color quenching, fluorescence quenching, and autofluorescence from the compound library. However, collisional quenching between the test compound and long fluorescence lifetime lanthanide chelates continues to be a problem for the homogeneous time-resolved fluorescence method. To overcome this problem, the DELFIA method [44], in which separation and washing steps are included, can be used as an alternate, but is not well suited to miniaturized formats because of the need for wash steps. All of these fluorescence-based techniques require labeling of reagents (substrate, ligand, and protein) with fluorophores. To reduce responses to unintended factors, temperature and pH-insensitive fluorophores with long-emission wavelengths are preferred. Since the labeling procedure may be time- and resource-consuming, the uses of generic reagents, such as fluorescent conjugates of streptavidin, GST, and IgG, are preferred. The label itself, or the labeling procedure, may alter the properties of the molecular interaction; therefore, labeled reagents should be subjected to thorough pharmacological characterization prior to use. Purified targets are generally preferred for biochemical assays, however, for those difficult to purify proteins, crude or partially purified cell extracts may be used. To maintain high-quality assay performance, thorough target characterization, in terms of sequence, pharmacological activity, stability, specificity of activity, and batch to batch variation, is mandatory. 15.4.2

Cell-Based Assays

Cell-based assays mimic the biological role of the target in a specific disease state more closely than non-cell-based assays, and have increasingly been used in HTS. At times they are the only choice for targets

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which require the whole cell for its function. Cell based assays used for the study of enzymes, receptors, transporters, and ion channels have been reported. Many of them are fluorescence or luminescence based, and miniaturizable [45,46]. The most commonly used cell-based assay is the ‘‘reporter gene’’ system [47,48]. If a gene is known to be activated as a result of a specific stimulus, the promoter region along with upstream enhancer elements may be fused to a readily measurable ‘‘reporter.’’ In response to a desired stimulus, the cell will produce the reporter. Normally, reporters are enzymes with luminescence substrates, such as luciferase, or with fluorescence substrates, such as betagalatosidase and beta-lactamase. The beta-lactamase reporter gene method can be configured into a FRET readout using a novel substrate that changes emission wavelength upon the cleavage of the beta-lactam ring [49]. Green fluorescence protein (GFP) and its various mutants have also been applied in cell-based assays to construct FRET and fluorescence imaging readouts. For protein-protein interaction, partners of the interaction pair are fused with different mutants of GFP [50]. For calcium mobilization, two types of GFPs are linked to a calmodulin and changes in the structure of calmodulin upon binding calcium bring the GFPs together or apart [51]. For signal transduction of orphan Gprotein-coupled receptors, trafficking of GFP tagged arrestin can be visualized using fluorescence imaging systems [52]. Technology capable of simultaneous sample injection and detection has made miniaturized HTS possible for calcium-sensing and membrane potential-sensing dye-based assays, where transient and weak fluorescence pulses occur seconds to minutes after stimulation by the ligand. The cell-permeable, calcium-sensing dyes have been widely used to study signal transduction systems where the activity of a membrane receptor is coupled with calcium mobilization. Membrane potential-sensing dyes have been used to study ion channel activity. An innovative FRET-based approach has been reported [53]. Miniaturized cell-based assays have been reported for suspension cells using a luciferase reporter [27]. The development of a miniaturized assay for adherent cells is a challenge due to the difficulty in maintaining the vitality of the cell under compact conditions. Advancements in cell culture and molecular biology techniques will benefit future assay development for adherent cell assays. To ensure high quality for miniaturized cell based assays, stable

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transfected cell lines are preferred over transient types. Other important factors to be considered are the level of endogeneous targets produced by different types of cell lines, the stability of target expression over repeated cell passages, the identification of cytotoxic compounds, protein synthesis inhibitors, and DNA binding compounds. There are unique problems associated with both biochemical and cell based miniaturized assays aside from liquid handling and signal detection. First, as the reaction volume decreases, the surface to bulk ratio increases. This will accelerate the evaporation of reagents, amplify background signals caused by reagent adsorption to the well surface, and increase light scattering and reflection by the plate. Second, as a homogeneous assay, high background caused by nonspecific binding between the target and the fluorophore is frequently encounted. Humidified chambers and hydrophilic reagents such as glycerol have been successfully applied to address the reagent evaporation issue [43]. Shortened reaction incubation time is an alternate for reducing evaporation. The use of solid black plates will minimize the light scattering. Finally, surfactants that can reduce nonspecific binding between reagents or between reagent and surface without denaturing the target protein, can be used as additives in the reaction solution or pre-coated onto the plate surface. 15.5 DATA HANDLING FOR MINIATURIZED ULTRA-HTS Until recently, the 96-well microtiter plate was the standard format for HTS, and the results were obtained in the 8 ⫻ 12 format for all assays. The drive toward miniaturization has given rise to a plethora of plate configurations, from 384 through 9600 wells. Thus, data capture and processing has to be sufficiently flexible to accommodate a variety of formats, usually consisting of multiples of 96. Most software for HTS allows one to define a template that mimics the plate layout. Provided there are definitions for compound identifiers in the desired format, the only limitations on density are the field space in the database (usually running on a remote server) and the ‘‘front-end’’ software that captures the data and presents them to the user. Import definitions can be adjusted to match output from the detection instruments, so that list or

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array formats for a variety of plate configurations can be accommodated. So-called prerefinement calculations, such as determination of ratios of fluorescence counts at different wavelengths, are usually handled by the instrumentation, but can also be performed by the HTS software by using a suitable data definition template. Thus, it is feasible to accommodate data derived from the novel detection methods that are essential for miniaturized formats. It is likely that the HTS community will move toward standardization of plate types [54], and particular plate configurations will be preferred. Currently, the 1536 layout (32 ⫻ 48 wells) is the most prevalent format for miniaturized assays. This allows mapping of compound identifiers from a storage format (usually 8 ⫻ 12 or 16 ⫻ 24) to the assay format in a relatively straightforward manner. A key requirement for all data handling systems, but particularly for miniaturized formats, is the implementation of a robust tracking system for compound identifiers. The system must be able to track the path of a compound as it is aliquoted and transferred from one container to another. The history of each compound must be traceable to its origin and synthetic procedure, and storage conditions should be captured for each replicate. This requires a robust interface between the database that stores chemical library information and the pipettors, barcoders and shipping contractors that perform day to day operations on the compound plates. For miniaturized systems, there are format changes that must be mapped back to the storage plates, and the data handling system must be capable of associating multiple source objects with each destination (assay) plate. If the organization has multiple sites, a global system is necessary for compound registration and tracking and procedures must be standardized. The identifier for each compound should reflect its history with respect to batch, storage conditions and replicate as well as its structure. 15.5.1

Quality Control

Since miniaturized assays are much more sensitive than conventional formats to minor variations in instrumentation and plate tolerances, effective quality control is important. Approaches to monitoring of data quality for miniaturized assays are generally similar to those taken for

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conventional formats [55,56]. Each plate has a set of controls (activated, no effect) and if available, a standard (known inhibitor or activator) in a predefined location. At a minimum, these controls must be monitored on a plate by plate basis to ensure consistency, usually defined as results within 3 SD of the validation controls. Additionally, it is desirable to monitor the standard deviation of inactive compounds [55] as a measure of ‘‘noise’’ throughout the screening campaign. Current liquid-handling equipment for miniaturized assays is not capable of simultaneous transfers to or from all wells in a 1536 or higher density plate. Reagents are added to groups of either 16 ⫻ 96 or 4 ⫻ 384 wells. This can create positional effects due to time-dependent changes in the properties of the reagents or to carryover from one group to another. Software for detecting such effects is available [57] and can also be used to apply corrective ‘‘normalization’’ to the data if the positional effects cannot be avoided by other means. 15.5.2

Data Volume

One of the aims of miniaturization is to increase the number of compounds that can be tested. In most cases, a library size of at least 1 million compounds is envisaged, with a throughput of 100,000 data points per day. This translates into about 3 megabytes of data per assay per day, or 30 megabytes per screen, a relatively modest amount. Performance of the hardware is unlikely to be limiting, unless multiple screens are being run or analyzed simultaneously. Problems can sometimes arise where data are being transferred across a network, or servers are being shared with other functions. Such problems can usually be solved by the use of a dedicated server for HTS applications. This becomes more important when data mining and visualization require simultaneous access to data from multiple screens, as well as chemical structure information. Current hardware and software are adequate for such applications, but it is important to plan ahead and ensure that the capacity and performance of the data handling system is matched with expected growth in library size, numbers of assay targets and HTS throughput. Increasing the throughput also places indirect demands on the infrastructure required for compound tracking and hit plate preparation (‘‘cherry picking’’). Seamless data transfer between HTS and com-

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pound supply, including automated storage and retrieval systems, is essential to prevent bottlenecks that could undermine the increases in throughput gained via miniaturization. 15.5.3

Visualization and Data Mining

Miniaturization is expected to yield large amounts of data, in terms of both numbers of measurements per assay and numbers of assays. Despite the ability of current computer systems to handle such data, human intervention is ultimately required to reach decisions. Hit detection can be done purely on a statistical basis, and the resulting list passed to the storage and retrieval software for cherry picking. Concentrationresponse profiling can also be automated, but as the number of compounds increases, it becomes impractical to visually examine every graph, and an automated procedure for quality assurance is necessary. Statistical analysis can be used to assess each concentration-response profile for goodness of fit, agreement between replicates, departure of high and low bounds from controls and excessively steep or shallow slopes. This can be automated and used to group compounds according to the quality of their profiles. Structural purity checking is often done at this stage, so the end result of the process is a ‘SAR list’ comprising biological activity with statistical ranking, and chemical structures with purity criteria, all of which can be done without human review. The ideal for a data handling system that supports miniaturization programs is to automate as much as possible of the decision making, by use of statistical procedures, and only require human review of the exceptions. The next step is to select compounds for follow-up, based on their biological and chemical properties. Currently, this requires human intervention, which may become impractical for very large screening campaigns. For biological activity, concentration-response data in the current versus historical screens are used to select compounds that show the desired potency and selectivity. Graphical representations offer the best approach to comprehension of large data sets, and software for such displays is available [58]. Chemical structures can be sorted according to molecular weight, absence of undesirable substructures (e.g., Michael acceptors, alkylators, etc.), ease of synthesis, patentability, purity, and clogP. In principle, these analyses can be automated,

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but the decision to accept or reject compounds depends on multiple factors and some degree of value judgement is usually required. Advances in computational chemistry and a better understanding of what constitutes a ‘‘druglike’’ molecule are needed to facilitate such decision making for very large data sets. 15.5.4

Hit Follow-up

A further consequence of miniaturization is the need for improvements in the throughput of secondary assays for hit follow-up. This may be mitigated somewhat by data mining approaches as described above, but increases in the numbers of compounds screened may for some assays give rise to large numbers of hits, requiring high throughput secondary assays. In turn, this requires both vertical (within assay) and horizontal (between assay) data analysis. A number of companies are pursuing early ADMET (absorption, distribution, metabolism, elimination and toxicity) assays in high-throughput formats. Data from these assays can be analyzed along with potency and secondary assay results by using multivariate approaches [59], which not only provide information about the effects of structural changes on individual factors, but also about interactive effects on multiple factors. This is ideally suited toward ‘‘lead explosion’’ techniques, where combinatorial chemistry or automated high speed parallel synthesis are used to prepare libraries around hit structures from HTS. Provided that the secondary assays are capable of testing libraries in the 1000–10,000 component range, compounds can be selected by clustering in the multidimensional space defined by the secondary and ADMET assays and by the target product profile. This approach requires extensive automation of the hit followup assays and is part of the ‘discovery factory concept’. 15.6 APPLICATION OF BIOCHIP TECHNOLOGY TO HTS Biochip technology is a new class of analytical tool in which a biological recognition event is integrated directly with signal discrimination, transduction, amplification, and detection. A classic example is the DNA-based chip technology where DNA arrays are immobilized to the

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surface of glass, plastic, quartz, or silicon by lithographic methods. In this instance, DNA hybridizations are analyzed by fluorescence-based technologies. High-throughput screening programs relating to drug discovery are increasingly embracing biochip technology; yet, they continue to lag behind the DNA-based chip technologies. This is in part due to the fact that in such an application for HTS, one would have to deliver the targets to the reaction site without compromising biological activity, and is also due to difficulties in delivering compounds from storage to the chip. The key difference between chip-based assays for DNA and for HTS applications is that the former makes multiple measurements on one sample while the latter makes one measurement on multiple samples. Because of such stringent conditions, which do not necessarily apply to the DNA-based technologies, the development of chip-based analysis for HTS remains a major challenge. Recently, however, innovative methods have led to the development of protein modified chip surfaces which include direct capture of the target on the surface, and transport of the target ‘‘in situ’’ by means of controlled microfluidic techniques. One technique to directly capture the target uses chemical linker arrays [60]. These arrays, such as silanes with the appropriate functional groups, are first coated on the chip surface using conventional photoresist lithography, photochemistry, and self-assembled monolayers. Following the activation of the functional group on the chip surface, target proteins or cells can then bind to the activated functional groups through their own surface reactive groups, or through affinity capture, such as antigen-antibody or strepavidin-biotin interactions. Another direct capture approach consists of constructing lipid bilayers on the chip surface. The target protein is then intercalated or conjugated in the membrane [61,62]. Advances in micromachining and microfluidics have facilitated the transport of target protein and other reagents to reaction sites synchronously without target immobilization at nanoliter or even picoliter levels with high precision. The delivery pathways are microscopic channel networks etched into the chip surface or inside the chip [63–66]. The migration of the reagents can be controlled without mechanical valves and pumps by establishing a voltage across various portions of the chip. The microscopic channel delivery method normally preserves the biological activity of reagents. In order to transduce the biological recognition events into detect-

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able physical phenomena, various methods have been examined, including electrochemical, optical, piezoelectric and thermometric transducers. For example, the conductance of molecular ion channels embedded in surface bound membranes can be switched by the molecular recognition event for digoxin and its antibody linked to the channels [62]. When the interaction partners are labeled with the appropriate donor and acceptor fluorophores, the molecular interaction between cholera toxin receptor and ligand taking place on the chip surface can induce distance-dependent FRET [61]. The refractive index at the chip surface changes as receptor binds to immobilized ligands [67]. An integrated biochip does not only contains biological recognition and signal transduction units, but could also bear discriminators. For examples, an electrical field can be applied to enrich charged reagents such as DNA, cells or proteins, to specific reaction sites, and bring products such as peptides, to desirable detecting positions [63]. Electrophoretic units can be incorporated to achieve rapid, high-resolution product separation for kinase and protease assays [64–66]. The miniaturized, versatile biochip technology could provide a viable route for screening larger number of compounds than currently feasible by the conventional microtiter plate based system. This approach would lower the cost of the reagents, while at the same time accelerate drug discovery efforts. However, the microfluidic reagent delivering system for biochips is not compatible with the current compound library infrastructure, which is based on microtiter plate formats. Therefore, compound access becomes the major obstacle to apply biochip technology in HTS. Practical uses of the biochip technology in HTS could start with combinatorial library screening where both parallel chemical synthesis and compound screening take place on the same chip [66]. Other uses of the technology in the near future can be lead optimization and toxicological studies. However, the compatibility between biochip technology and compound library infrastructure remains to be solved. 15.7 EXAMPLES OF MINIATURIZED HTS ASSAYS Reports of assay validation experiments performed in 1536 plates are becoming commonplace, and presentations at HTS conferences have

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documented demonstration of feasibility for almost every type of common HTS assay in the 1536 format. Notable exceptions to this generality are cell based kinetic membrane transport assays (currently possible in 384 well plates), and filtration type assays (only possible in 96-well plates at this time). To date, however, there have been very few reports of full implementation of miniaturized truly high throughout screens (defined as batch screening of a compound collection ⬎25,000 samples on a target). Two 1536-well, 40,000-compound assays, a fluorescent proteolytic and a suspension cell-based, have been described [13]. Several reports of validation of fluorescence reported enzymatic assays have appeared [10,11,27,68–71]. Validation of an absorbance-based proteolytic assay using a CCD imager was reported. Chemiluminescent and chromogenic implementations of a washed 1536-well immunoenzyme amplified tyrosine kinase assay using a CCD imager have also been reported. A thorough comparison of 96- 384- and 1536-well formats to screen a colorimetric whole cell transcriptional assay appeared in 1998 [73]. The challenges posed by distributing cells evenly into plates, culturing them for a limited amount of time, lysing the cells and detecting luminescence have been overcome. Chemiluminescent suspension cell-based transcriptional assays have been reported by several groups [12,27,68,69]. To date, there have been no reports of validation of adherent cell assays in miniaturized format. 15.7.1

Mass-Dependent Assay: Fluorescence Polarization (FP)

A novel assay for measuring the activity of adenine transferase that transfers multiple adenine-containing groups to an acceptor protein has been developed using fluorescence polarization technology in 1536well plate format [43]. In the assay, a long wavelength fluorescence tracer was covalently conjugated to the adenine of the donor substrate through a C 6 spacer arm. As a result of the transfer of the adeninecontaining moieties to the acceptor protein substrate, the rotation correlation time of the fluorescence tracer increased, and hence the degree of fluorescence polarization. The pharmacological profile and kinetics of the enzyme measured according to the fluorescence polarization method were consistent with those determined previously by a conven-

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Table 1 Assessment of the Performance of the Fluorescence Polarization Assay for Adenine Transferase by Statistical Analysis

Mean Value (mP) Standard deviation (mP) Assay value ratio* (AVR)

Signal

Background

154.1 10.5

25.8 9.2 0.46

Standard compound 61.0 11.9

3(SD signal ⫹ SD background ) MEAN signal ⫺ MEAN background An AVR value of 0.6 or less is a prerequisite for effective HTS. Source: Ref. 55. *AVR ⫽

tional filtration-based method. The assay was successfully applied to execute a 250,000-compound high-throughput screening program. Table 1 summarizes the performance of the assay by statistical analysis. A fluorescence polarization based assay was designed for screening competitive inhibitors of a nuclear receptor. In this assay, the degree of fluorescence polarization of a fluorescein-labeled ligand increases as the ligand binds to the receptor. The assay was miniaturized from the original 100 µL in a 96-well plate format to 30 µL in a 384-well plateformat, which was further miniaturized to 4 µL in a 1536-well plate format. The IC 50 value of a known competitive inhibitor of the receptor obtained from the 1536-well plate assay is consistent with those obtained from the 96-well and 384-well plate assays, as shown in Figure 4. 15.7.2

Proximity-Dependent Assay: Fluorescence Resonance Energy Transfer (FRET)

The peptide substrate of a protease consisting of 11 amino acids was labeled with a Texas Red and a Cy5 fluorophores at the N-terminus and a lysine that is 7 residues apart, respectively. The Cy5 kept the Texas Red moiety quenched until the protease clipped the substrate between the two dyes, thereby releasing the Texas Red which was excited at wavelength 580 nm and emitted at wavelength 620 nm (Fig. 3). The pharmacological and kinetic characteristics of the enzyme ob-

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Figure 4 Inhibition of the ligand-receptor binding by a known inhibitor in 96-, 384-, and 1536-well plate formats. The degree of inhibition was monitored by the fluorescence polarization changes of a fluorescein-labeled ligand.

tained by the fluorescence resonance energy transfer method were consistent with those determined by fluorescence-quenching methods using coumarin (λem ⫽ 420 nm) containing substrates. The false positive rate obtained from the FRET based assay method was significantly lower than that from the later method, presumably due to the elimination of interferences of light absorbance, emission and scattering from testing compounds and labware, since these interferences peak at a range from yellow to green wavelengths. Figure 5 shows the kinetic measurement of the assay in 1536-well plate format. 15.7.3

Luminescence-Based Assay

The luciferase reporter gene system is frequently used for cell based transcription assays. The 1535-well plate formated assay was designed for screening inhibitors of purified firefly luciferase. In the presence of ATP, the luciferase oxidizes luciferin and results in luminescence. To overcome the weak and unstable signal problem associated with the luminescence based assay, a highly sensitive CCD camera was used

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Figure 5 Measurement of protease activity by fluorescence resonance energy transfer method in 1536-well plate format. The substrate for the protease was labeled with Texas Red and Cy5 dyes. The kinetics of substrate cleavage by the enzyme was monitored by the Texas Red fluorescence intensity changes.

for detection. Figure 6 gives the test results sorted by the degree of inhibition. This diagram demonstrates a relatively clear differentiation of the hits from the noise of the assay. Greater than 85% of hits previously identified by the 96-well plate formatted method were confirmed by the miniaturized method. 15.7.4

Bead-Staining Assays

Combinatorial libraries synthesized as one compound per bead form the basis of one of the earliest reported miniaturized screening methods. In the affinity-based staining technique, up to 1 million compounds can be screened for target binding in a volume of 10 mL by one technician in ⬍4 hr. Although limited to a small subset of targets, this method achieves extremely high throughput and very low effective screening volumes (10 nL per compound assayed), but can suffer from artifacts

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Figure 6 Statistical distribution of data generated by the inhibition of 3236 compounds in a luminescence-based luciferase assay. The slightly distorted Gaussian curve is characterized by a mean inhibition of ⫺1.4% and a standard deviation of ⫾11.4%.

owing to the very high local concentration of compound in the bead microenvironment. 15.7.5

Agar-Embedded Bead Assays

A free-format miniaturized screening assay for combinatorial libraries that are synthesized on beads has been reported. In this approach, compounds are released from beads just prior to dispersion in agar, and activity is detected in the area of diffusion around each bead. This method allows very high throughput, but requires the use of special photocleavable linkers and an imaging system for automated detection and quantitation of compound activities.

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15.8 ISSUES FOR MINIATURIZATION 15.8.1

How Low Is Low Enough?

Current fluorescence techniques are capable of detecting single molecule events [74–76]. In principle, one could pursue miniaturization to this ultimate level. However, this is not necessarily desirable, either from a practical or theoretical standpoint. Populations of protein molecules may differ according to their degree of folding, and in a ‘quantized’ assay, these changes could have an all-or-none effect on function. Provided that the source population of protein molecules is large compared with the number of data points in the assay and that it is normally distributed with respect to folding, one would expect sampled individual molecules to be representative of the average population. However, if the protein is impure or contains grossly misfolded molecules, one could select non-functional molecules for the assay. Small changes in the microenvironment of the target molecule could also lead to excessive variability across a replicated single molecule array, resulting in poor signal to noise ratios. In bulk populations of molecules, these extraneous effects are averaged, whereas single molecules may be affected either by the sensitivity of the molecule to the effect or by the magnitude of the effect itself. The latter is unpredictable, and depends on the assay configuration. With proper attention to the purity and specific activity of the target protein, and to homogeneity within and between assay compartments, single molecule screening could theoretically be accomplished for simple assays using purified proteins. For cell-based and membrane-bound targets, however, where the minimum ‘‘unit’’ of activity is one cell or one membrane vesicle, variability between cells or vesicle properties (size, density of protein molecules, purity) is limiting. For cell-based assays, the minimum number of cells for representative signal generation has been empirically estimated as 100. For a cell volume of 5 pL, the minimum theoretical assay volume is 500 pL packed cell volume plus the volume of medium necessary to sustain the cells for the duration of the experiment. Practical considerations are the primary limitation when deciding how far to miniaturize assays. Issues with mixing of reagents, evaporation, droplet formation, positional accuracy of instrumentation, cost,

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reformatting of compound libraries, complexity of assay development, plate tolerances, surface adsorption, and customer needs require the assay designer to strike a balance between miniaturized and conventional formats. As technology improves, some of these limitations will undoubtedly become less significant, but the degree to which an organization is prepared to invest in such technologies will ultimately be driven by need. Library size and reagent conservation are the major drivers of miniaturization technology. There is a growing realization that simply increasing the size of the library does not linearly increase the number of drug candidates. The ability to find active compounds is more dependent on the target type than on the number of compounds tested. For ‘‘classical’’ targets that have small and deep binding pockets, such as G-protein-coupled receptors or enzymes, testing of relatively small numbers of compounds has yielded active agents, and the main benefit of larger libraries is to provide more structural classes to reduce attrition in the later phases of lead optimization. For proteinprotein targets, where interactions are dispersed across a large area and solvent exclusion contributes to binding, it is unlikely that quantitative changes in the library will improve hit rates, and qualitative changes in the type of compound and/or strategy for drug discovery are required. Thus it is likely that in future, more attention will be given to the content rather than the size of the library. This may be expected to influence investment decisions away from further miniaturization and in favor of computational chemistry, data mining, and novel synthetic approaches. 15.8.2

Timing

If the sole reason for assay miniaturization is to enable a cost efficient large scaleup in screening capacity, why is there a sense of urgency to miniaturize? Reliable 1536 equipment is only now beginning to come to market, and the technology will undoubtedly mature and become much more robust in the coming years. What is the cost of waiting to invest until the equipment is ‘‘turnkey’’? We are at a crucial point in the history of the HTS and the pharmaceutical industry as a whole. All of the industry leading pharmaceutical companies have committed to the goal of introducing two to three significant new chemical entities

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per year, and most have acknowledged that this leads to the need to screen 75 to 100 targets per year. Current estimates from the Human Genome Project are that there are ⬃30,000 human genes, and a significant portion of those genes are estimated to be useful targets for therapeutic intervention. If the top 10 pharmaceutical companies were to screen 100 new, unique targets per year, projects for each of the human therapeutic targets will be under way within 10 years. In practice, pharmaceutical companies tend to focus on targets within the same gene families, so the majority of novel approachable targets will be spoken for much earlier than this. This implies that there is great urgency to begin uHTS, and the only cost-effective route is via assay miniaturization. 15.8.3

Productivity Measurement

How does one measure the benefits or detriments of miniaturization? Reduction in the assay volume is expected to lead to savings in cost associated with reagent production, greater throughput, and reduced consumption of library compounds. However, this is offset by greater capital equipment costs, longer cycle times for assay development and troubleshooting, increased down time associated with use of unproven technologies, and increased cost of consumables such as plates and pipette tips. The total cost of a miniaturized assay, including FTEs, consumables and capital can be compared with equivalent costs for a conventional assay. Some figures are given in Table 2. Throughput and cycle time are not necessarily equivalent owing factors associated with assay development and down time. Time from reagent receipt to delivery of concentration-response data to customers is a measure of all phases of the screening campaign and can be used to evaluate the effects of miniaturization on cycle times. Throughput can be determined as the number of data points generated per 24-hr period (averaged over the duration of the screen). A widely used measurable for screening campaigns is the number of compounds taken up by the medicinal chemistry group. It is important to show customers that miniaturization does not result in failure to detect all the compounds that would be found in a conventional screening format. Finally, the proportion of target types that can be accommodated in a miniaturized format is also

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Table 2 Cost Comparison of Conventional and Miniaturized Assay Formats for an Enzyme Assay with 200,000 Compounds

Item Reagents Consumables Waste disposal Capital FTE costs Total

Conventional Format: Radioactive Readout in 96-well Plates ($)

Miniaturized Format: Fluorescence Readout in 1536-well Plates ($)

490,000 82,000 90,000 206,000 84,000 952,000

10,000 2,000 1,500 370,000 21,000 404,000

an important indicator of the scope of the technology and of the need for additional development. Clearly, these are moving targets as technology changes, and many companies opt to maintain some form of miniaturization program simply to keep up with the latest developments. However, the expectation is that these investments will lead to measurable improvements in deliverables as above. 15.8.4

Education of Customers

In most organizations, biological targets are identified in one group and high-throughput screening is done in another. In some cases, a separate ‘‘technology development’’ group is established to pursue novel approaches to assay development and HTS. This can lead to ‘‘technology gaps’’ between groups such that assays are developed using radioactive or other nonminiaturizable approaches, and time-consuming reformatting is needed when the assay is transferred. Additionally, skepticism about novel assay configurations and the perception that the technology group is merely ‘‘playing’’ with expensive equipment can negatively impact efforts toward assay miniaturization. The origin and solution to these problems lies in the ability of the groups to communicate clearly with one another. Mentoring approaches in which individuals from the HTS and/or technology groups participate in projects from their inception, can redirect assay development efforts toward miniaturizationcompatible formats. Temporary assignments of individuals between

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groups can provide additional perspective, and seminars describing real-life examples of benefits derived from miniaturization can help dispel skepticism and promote enthusiasm for novel approaches. Any novel assay procedure must always be tested with compounds of known activity, if available, so that there are no doubts about its ability to find bona fide inhibitors or activators. 15.8.5

Obsolescence Cycles

An inevitable consequence of the pace of technological development is the rapid obsolescence of hardware and software associated with high-throughput screening. A dilemma exists between the desire to have access to the latest technology and to contain costs associated with equipment turnover. Three approaches have been taken to solve this problem. The first is to buy instrumentation that is upgradable— i.e., components can be replaced without having to purchase a new instrument. This is effective where a robust basic platform is available, onto which components can rapidly be exchanged with minimum downtime. However, the platform itself can become obsolete or the vendor can go out of business, and the cost of upgrades can be significant. The second approach is to outsource all technology development, such that the vendor provides instrumentation or services under a partnership arrangement, in which obsolete products are replaced with the latest version. This approach can be particularly effective for software, which requires constant maintenance and user support. Additionally, companies that have patented technologies often demand partnerships as a precondition for access to their products or services, but such arrangements can be very expensive. Partnerships can be very helpful, possibly essential, where the technology is highly experimental, and the user does not wish to spend time fixing ‘‘bugs.’’ A disadvantage of the partnership approach is that the user is tied to one supplier and cannot ‘mix and match’ disparate products. A third approach is to lease instruments, either directly from the vendor, or via a third party. This is useful when a particular instrument is expected to have a short service life, at which point the lessor assumes responsibility for the instrument and the lessee is free either to buy or lease the newer version. The disadvantage of leasing is that one needs to predict the service life

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of the instrument, which is not always possible with new technologies. Most companies adopt a combination of these approaches, dependent on the mix of technologies that is desired. 15.8.6

Compound Handling

An often neglected aspect of high-throughput screening is the need to manage large chemical libraries. Compounds arriving into the collection must be registered, weighed, solubilized, formated into master plates, and redistributed into ‘‘daughter’’ plates for assays. Active compounds from screens must be ‘‘cherry-picked’’ into hit plates and distributed for concentration-response determination and structure confirmation. The ‘‘macro-to-nano’’ interface between a large compound store and a miniaturized assay format requires specialized instrumentation, capable of dispensing very small volumes of liquid. Problems can arise with poorly soluble or viscous compounds, cross-contamination, and mixing in small volumes. The ideal arrangement would be to aspirate from a macro scale master plate using a disposable probe, dispense directly into the miniaturized plate or microfluidic system, and return any unused material to the master plate. Volume capability should be such that the need for intermediate dilutions is eliminated. To date, there is no instrumentation that can accomplish this reliably. A compromise approach is to aspirate a moderate volume from the master plate into a daughter plate using pipettors with disposable tips, and use the daughter plate for several screens, after which it is discarded. Modest cross-contamination in the daughter plate can be tolerated, and solubility problems can be identified by turbidimetric analysis of the daughter plate. Some groups opt to dispense directly from the master plate into several assay plates and dry the compound for storage and subsequent resolubilization. This avoids intermediate steps, but many compounds will fail to dissolve after evaporation, thus the screening concentration may differ from that desired. 15.9 SUMMARY Developing robust miniaturized assays for HTS not only requires highly trained individuals with multifaceted scientific backgrounds, but

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also demands time and financial commitments. Nonetheless, the ultimate time, cost, and reagent savings for outweigh the developmental challenges, undoubtedly making assay miniaturization an important focus for HTS. REFERENCES 1. Rasvi, E.S. (1999). The drive to miniaturize. Drug Market Dev., 10: 211–214. 2. Burbaum, J.J. (1998). Miniaturization technologies in HTS: how fast, how small, how soon? DDT, 3:313–322. 3. Sills, M.A. (1998). Future considerations in HTS: the acute effect of chronic dilemmas. DDT, 3:304–312. 4. Wallace, R.W. (1998). A little too much? Expense versus return in HTS miniaturization. DDT. 3:299. 5. Knebel, G. (1998). High-throughput screening with 1536 wells. Am. Biotechnol. Lab. News, March 1998. 6. Fox, S., Farr-Jones, S., and Yund, M.A. (1999). High throughput screening for drug discovery: continually transitioning into new technology. J. Biomol. Screening, 4:183–186. 7. Rose, D. (1999). Microdispensing technologies in drug discovery. DDT, 4:411–419. 8. Brandt, D.W. (1997). Multiplex nanoliter transfers for high throughput drug screening using the Biomek 2000 and the high density replication tool. J. Biomol. Screening, 2:11–116. 9. Ramm, P. (1999). Imaging systems in assay screening. DDT, 4:401– 410. 10. Oldenburg, K., Zhang, J.H., Chen, T., Maffia, A., Blom, K.F., Combs, A.P., and Chung, T.D.Y. (1998). Assay miniaturization for ultra HTS of combinatorial and discrete compound libraries: a 9600-well (0.2 µL) assay system. J. Biomol. Screening, 3:55–62. 11. Abriola, L., Chin, M., Fuerst, P., Schweitzer R. and Sills, M. (1999). Digital Imaging as a detection method for a fluorescent protease assay in 96-well and miniaturized assay plate formats. J. Biomol. Screening, 4:121–127. 12. Vollert, H. (1998). Development of a robust miniaturized screening system. Proceedings, IBC, Practical Aspects for Assay Miniaturization and Design for Drug Discovery, Boston, MA.

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13. Vollert, H. (1999). HMR’s pilot screens in 1536. Proceedings, IBC, Miniaturization Technologies Conference, Berkeley, CA. 14. Vollert, H. (1999). Performance of a miniaturized screening system and future methods of detection. Proceedings, IBC, Drug Discovery Technologies Conference, Amsterdam, Netherlands. 15. Weiss, S. (1999). Fluorescence spectroscopy of single biomolecules. Science, 283:1676–1683. 16. Webb, W.W. (1976). Applications of fluorescence correlation spectroscopy. Q. Rev. Biophys. 9:49–68. 17. Pitschke, M., Prior, R., Haupt, M., and Riesner, D. (1998). Detection of single amyloid beta-protein aggregates in the cerebrospinal fluid of Alzheimer’s patients by fluorescence correlation spectroscopy. Nat. Med., 4:832–834. 18. Auer, M., Moore, K.J., Meyer-Almes, F.J., Guenther, R., Pope, A.J., and Stoeckli, K.A. (1998). Fluorescence correlation spectroscopy: lead discovery by miniaturized HTS. DDT, 3:457–465. 19. Winkler, T., Kettling, U., Koltermann, A., and Eigen, M. (1999). Confocal fluorescence coincidence analysis: an approach to ultra highthroughput screening. PNAS, 96:1373–1378. 20. Sto¨ckli, K. (1999). Confocal fluorescence techniques: implementation and applications in lead discovery and validation. Proceedings, IBC High-Throughput Screening Conference, Berkeley, CA. 21. Eigen, M., and Rigler, R. (1994). Sorting single molecules: application to diagnostics and evolutionary biotechnology. PNAS, 91:5740–5747. 22. Kettling, U., Koltermann, A., Schwille, P., and Eigen, M. (1998). Realtime enzyme kinetics monitored by dual color fluorescence cross-correlation spectroscopy. PNAS, 95:1421–1426. 23. England, P.J. (1999). Industrializing drug discovery: ultra highthroughput screening in 3,456 plates. IBC’s Miniaturization Technologies Conference, Berkeley, CA. 24. Springhorn, J.P., Rollins, S.A., and Squinto, S.P. (1999). Drug discovery and screening on a compact diskette. IBC’s Miniaturization Technologies Conference, Berkeley, CA. 25. Brandt, D.W. (1998). Core system model: understanding the impact of reliability on high throughput screening systems. DDT, 3:61–68. 26. Bosworth, N., and Towers, P. (1989). Scintillation proximity assay. Nature, 341:167–168. 27. Maffia, A.M. III, Kariv, I., and Oldenburg, K.R. (1999). Miniaturization of a mammalian cell-based assay: luciferase reporter gene readout in a 3 microliter 1536-well plate. J. Biomol. Screening, 4:137–142.

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28. Hill, D.C. (1998). Trends in development of high throughput screening technologies for rapid discovery of novel drug. Curr. Opin. Drug Discover Dev., 1:92–97. 29. Schullek, J.R., Butler, J.H., Ni, Z.J., Chen, D., and Yuan, Z. (1997). A high-density screening format for encoded combinatorial libraries: assay miniaturization and its application to enzymatic reactions. Anal. Biochem., 246:20–29. 30. Pope, A.J., Haupts, J., and Moore, K.J. (1998). Homogeneous fluorescence readouts for miniaturized high throughput screening: theory and practice. Proceedings, DDT, 4:350–362. 31. Seethala, R., and Menzel, R. (1997). A homogeneous, fluorescence polarization assay for Src-family tyrosine kinases. Anal. Biochem., 253: 210–218. 32. Fernandes, P.B. (1998). Technological advances in high-throughput screening. Curr. Opin. Chem. Biol., 2:596–603. 33. Rogers, M.V. (1997). Light on high-throughput screening: fluorescencebased assay technologies. DDT, 2:156–160. 34. Kolb, J.M., Yamanaka, G., and Manly, S.P. (1996). Use of a novel homogeneous fluorescent technology in high throughput screening. J. Biomol. Screening, 1:203–210. 35. Mellor, G.W., Burden, M.N., Preaudat, M., Joseph, Y., Cooksley, S.B., Ellis, J.H., and Banks M.N. (1998). Development of a CD28/CD86 (B72) binding assay for high throughput screening by homogeneous timeresolved fluorescence. J. Biomol. Screening, 3:91–99. 36. Kolb, A.J., Kaplita, P.V., Jayes, D.J., Park, Y-W., Pernell, C., Major, J.S., and Mathis, G. (1998). Tyrosine kinase assays adapted to homogeneous time-resolved fluorescence. DTT, 3:333–342. 37. Stenroos, K., Hurskainen, P., Eriksson, S., Hemmila, I. Blomberg, K., and Lindqvist, C. (1998). Homogeneous time-resolved IL-2IL-2Rα assay using fluorescence resonance energy transfer. Cytokine, 10:495– 499. 38. Youn, H.J., Terpetschnig, E., Szmacinski, H., and Lakowicz, J.R. (1995). Fluorescence energy transfer immunoassay based on a long-lifetime luminescent metal-ligand complex. Anal. Biochem., 232:24–30. 39. Szmacinski, H., and Lakowicz, J.R. (1995). Fluorescence lifetime-based sensing and imaging. Sensors Actuator, 29:16–24. 40. Stryer, L. (1978). Fluorescence energy transfer as a spectroscopic ruler. Annu. Rev. Biochem., 47:819–846. 41. Gryczynski, I., Wiczk, W., Johnson, M.L., Cheung, H.C., Wang, C.-K., and Lakowicz, J.R. (1988). Resolution of end-to-end distance distribu-

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screening. In: Principles and Practice of High Throughput Screening. Murray, K., ed. Blackwell Publications, Oxford, UK. In press. Zhang, J., Chung, T.D.Y., and Oldenburg, K.R. (1999). A simple statistical parameter for use in evaluation and validation of high throughput screening assays. J. Biomol. Screening, 4:67–73. Web site of Novel Science: www.NovelScience.com/Eng/eng bsp en.html. Interactive Data Visualization. User’s manual and user reference by Spotfire Inc. Boston, MA. Neter, J., Wasserman, W., and Kutner, M.H. (1985). Applied linear statistical models: regression, analysis of variance, and experimental designs. Irwin Publications, Homewood, IL. Blawas, A.S., and Reichert, W.M. (1998). Protein patterning. Biomaterials, 19:595–609. Song, X., Nolan, J., and Swanson, B.I. (1998). Optical biosensor based on fluorescence resonance energy transfer: ultrasensitive and specific detection of protein toxins. JACS, 120:11514–11515. Cornell, B.A., Braach-Maksvytis, V.L.B., King, L.G., Osman, P.D.J., Raguso, B., Wleczorek, L., and Pace, R.J. (1997). A biosensor that uses ion-channel switches. Nature, 387:580–583. Cheng, J., Sheldon, E.L., Wu, L., Uribe, A., Gerrue, L.O., Carrino, J., Heller, M.J., and O’Connell, J.P. (1998). Preparation and hybridization analysis of DNA/RNA from E. coli on microfabricated bioelectronic chips. Nat. Biotech., 16:541–544. Knapp, M. (1999). Near-term solutions for nanoscale experimentation based on lapchip technology. Proceedings, IBC’s Annual International Conference on Miniaturization Technologies, Berkeley, CA. Boone, T.D. (1999). Rapid multiplexed analyses on plastic microfluidics. Ibid. Heaney, P.J. (1999). Massively parallel microfluidic instrumentation for drug discovery and pharmacogenomics. Ibid. Jonsson, U., and Malmqvist, M. (1992). Real time biospecific interaction analysis. Adv. Biosensors, 2:291–336. Brown, D.M., (1999). Scaling to 1536: using fluorescent and luminescent detection. Proceedings, IBC’s Assay Miniaturization Conference, Berkeley, CA. Berg, M., Undisz, K., Thiericke, R., Grabely, S., Moore, T., and Posten, C. (1999). Comparative study on the feasibility of enzyme assays and cellular reporter gene assays in yeast in the 96-, 384-, and 1536-well format. Proceedings, Laboratory Automation Meeting, San Diego, CA.

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16 Integrated Microsystems in Clinical Chemistry Constraints and Consequences Michael Bell Beckman Coulter, Inc., Brea, California

16.1 INTRODUCTION AND SCOPE Clinical chemistry is the measurement of diagnostically or therapeutically relevant molecules in patient samples. This chapter will focus on analytes dissolved in biological fluids, particularly blood and its fractions. It will not deal with cells or cell contents. Clinical chemistry has a number of special requirements different from other applications of microfluidic devices. These include a wide range of analytes, high cost sensitivity, extreme analyte dynamic ranges, messy samples, and fast turnaround. These requirements impose constraints on the application of microsystems that have implications on their design and use. 16.2 BACKGROUND—BREADTH OF ANALYSIS Clinical chemistry is a large field with a broad collection of analytical methods. Dividing the field by results urgency helps to clarify the application of microsystems.

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The results of tests for critical care analytes immediately affect treatment decisions. The menu of testing associated with surgery and trauma is limited to blood gases, pH, electrolytes, and a few metabolites (glucose, creatinine, urea, etc.), but this limitation is technologically imposed. Other emergency tests include markers of myocardial infarction (myoglobin, creatine kinase MB, and troponin), drugs of abuse, pregnancy (chorionic gonadotropin [hCG]), selected therapeutic drugs (digoxin and theophylline), and certain infectious agents. Microscale sensors measure many of the higher concentration analytes [1–3]. Routine analytes constitute the bulk of clinical tests. ‘‘General chemistry’’ analytes are commonly divided from ‘‘special chemistry’’ analytes, but the distinction is one of history and technology, not of application. Routine analysis provides information for diagnostic or therapeutic decisions where time is not critical—a few hours or a day does not make much difference in treatment. Such delays are becoming less tolerable as costs accrue if some medical decisions are postponed. This is particularly true when patients are in hospital. Most general chemistry analytes are of relatively high concentration (nanomolar to 100 mmolar); many were originally measured by colorimetric reactions that date back to the early 1900s. These have largely been replaced by assays based on biological reagents— enzymes and antibodies. These analytes have long been assayed in large panels as part of diagnostic fishing expeditions for aberrant values. This is hard to justify economically; where once 20–30 tests per sample was common, the average today is ⬍10 and declining. Special chemistry analytes are of lower concentrations (below nanomolar) and are measured by heterogeneous immunoassay or DNA binding. Most analytes are hormones, antibodies, and infectious agents. As techniques improve, more of these analytes are measured by homogeneous methods that are less costly to automate. Heterogeneous assays involve a separation step that permits removal of unreacted materials. The extra processing steps complicate automation, but improve the limits of detection. Hospitals use ‘‘stat’’ to designate samples for priority analysis. The turnaround time for stat assays depends on the analyte and on the conventions of the hospital, but one-half hour is a common target. This is the total time to deliver assays results—only a small fraction is used

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in the actual analysis. Most stat designations reflect diagnostic or therapeutic urgency, but the practice may be used to rule out conditions and free up resources. Some critical care analytes are processed as stats with target turnaround time of a few minutes. Most hospital analyzers have the ability to interrupt queued samples in progress to handle priority samples. The most common detection method for clinical chemistry analytes on conventional systems is spectrophotometry. Others include selective electrodes, fluorometry, and fluorescence polarization. There are some difficulties in applying spectrophotometry to microdevices due to the small path lengths, but similar measurements have been made on other small path length systems such as capillary electrophoresis and HPLC. These have also been demonstrated on microfabricated devices [4]. In addition, nearly all clinical chemistry analytes have been demonstrated by multiple methods, including fluorescence, which is well matched to microscale devices. 16.3 SPECIAL ISSUES Clinical chemistry has some additional issues as a consequence of the nature of its analytes and history. These include assay calibration, temperature control, and shelf life. 16.3.1

Precision, Accuracy, and Calibration

Clinical chemistry tests rarely run as absolute assays. Most measure a reaction and relate the measured value to analyte concentration by calibration. Running known value calibrators under similar conditions can correct for a multitude of errors. Repeatable conditions between sample and calibrator assays are essential to this scheme. Serum enzyme determinations are usually run without calibration. International standards [5–7] dictate measurement under carefully controlled conditions, particularly temperature. 16.3.2

Temperature Control

Most clinical chemistry systems control the temperature of reaction. In general, assays relying upon enzymes are more temperature sensitive

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and antibodies less so. Some systems run assays at measured ambient temperatures and correct the results by stored temperature sensitivity factors. This is practical when the sensitivities are known; many change with different manufacturing lots and must be empirically determined when reagents are manufactured. Serum enzyme determinations are perhaps an exception. The temperature-sensitive component is the analyte itself. Although population studies of enzyme temperature sensitivities can be run, individual samples may contain variant enzymes that do not match the population average. Despite this, it has become accepted practice to adjust enzyme activities measured at one temperature to effective values at a second temperature. The usefulness of this depends on carefully controlled measurement methods [8]. Microfluidic systems have excellent geometry for precise temperature control. Most are flat for reasonable contact with temperature control devices such as thermoelectric coolers. The content of channels forms a small fraction of the overall heat capacity of the device. Also, the thermal path between the walls and all elements of the fluid contents can be made very short. The result of this is that even if samples or reagents are far from the operating temperature when introduced, they can reach that temperature in a matter of moments without fastresponse control circuitry. This is an improvement over conventional-scale systems where the reaction mixture has a substantial fraction of the total heat capacity. Adding chilled reagents perturbs the system and requires thermal equilibration times up to several minutes. The longer thermal transfer distances further slow the process at the larger scale. 16.3.3

Shelf Life

Devices to run clinical chemistry tests must be available at the time of need. The practical consequence of this, given the vagaries of shipping and distribution networks worldwide, is that the units must remain active for at least 1, and preferably 2 years, after production. Ambient storage is preferable to refrigeration, but ambient shipping may subject the devices to temperatures in excess of 40°C for several days. Tiny volumes of preloaded reagents in a microdevice shift position during shipping. Liquids evaporate and recondense onto packaging

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materials, getting lost prior to use. Including additional geometry such as mechanical valves to isolate reagents can prevent these effects, but this increases device complexity. 16.4 COST CONSTRAINTS The single greatest constraint on clinical chemistry testing is cost. While selected new or esoteric analyses may be reimbursed to the health care provider at tens or even hundreds of dollars each, for the vast majority, diagnostics manufacturers receive ⬍U.S. $1.00; in most instances this amount is ⬍$0.20. The aggregate amount paid for clinical testing is unlikely to increase because the market is governed by noneconomic factors—in most cases neither the patient nor the ordering physician directly pays the costs. Third-party payers, governments, and insurers (in the United States) pay for the majority of testing. Aging populations increase the demand for clinical testing and other healthcare services, but the same demographic trends decrease the proportion of active workers and the amount of economic output available for health care expenses. Without continued substantial gains in productivity or politically unpalatable cutbacks in other areas, healthcare spending cannot increase in proportion to demographic demands. Clinical laboratory testing is one component of outcomes management studies wherein hospitals and third-party payers codify treatment options; each medical procedure is evaluated for costs and benefits. Selected testing may be increased to save costs in other, non-testing areas. The much more common result of such studies to date is that testing is reduced, eliminated, or staged with other procedures. These macro trends conspire to reduce, or at best maintain, the reimbursable costs of clinical laboratory testing. Any new technology aimed at other than small niches in clinical testing needs to pay particular attention to costs. A new technique that is more expensive than established methods will face high resistance. 16.4.1

Cost Limits of Microsystems

Microfabricated devices appear an ideal fit to this low cost requirement. Similar manufacturing techniques have reduced the costs of computing

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hardware 100,000-fold since 1970. These declines are primarily attributable to scaling and improved processing. Costs have declined per unit of function for large-scale integrated circuits (LSI), but complexity has increased to keep sales-unit prices high. Prices for small units of fixed functionality, such as single packaged gates, have declined much less. Their price is no longer controlled by the cost of the semiconductors, but by the cost of secondary operations and packaging. For example, a quad NAND gate cost $0.30 in 1970, but cost $0.38 in 1999 [9]. When adjusted for inflation, this translates into only a 3.4-fold decrease in price over 30 years. Breadboard integrated microfluidic devices today are far from the cost of a NAND gate, and it would be premature to speculate too precisely on their eventual production cost. However, single use devices share with the simple package of gates a floor to their cost—that of their packaging. Even if the microdevice and all of its reagent contents were free, it would still be subject to manufacturer standard costs of packaging. Table 1 compares the manufacturer packaging costs (in U.S.$) for a single use device and for a conventional 200-dose clinical chemistry kit. The cost for external packaging for a single use device ($0.26) exceeds the median manufacturer price for a single test. This means that such devices need to be targeted at niche applications where higher prices may be justifiable, or that the devices must be made to deliver more than one test result. 16.4.2

Device Reuse

A device may deliver more than one test result by profiling several assays for a single sample. This is limited by the health care outcomes management studies that have consistently reduced the number of tests ordered on a single sample at one time. In view of this, and in light of historical billing practices, third-party payers in the United States have capped the amount they reimburse for multitest profiles and will not reimburse when tests ordered do not correspond to the suspected diagnosis. A more palatable alternative is to use a microdevice to run tests on many different specimens. This may be accomplished by including

Single use

Item Foil pouch Unit label Pouch label Box Insert Shipping container Total

Multiuse

Quantity

Unit cost

1 1 1 1/20 1/20 1/200

0.10 0.05 0.05 0.60 0.60 1.00

Cost per result 0.10 0.05 0.05 0.03 0.03 0.005 0.265

Item Bottle Caps Labels Box Insert Shipping container

Quantity per kit

Unit cost

Cost per kit

Cost per result

1 3 2 1/2 1/2 1/10 Total

1.00 0.05 0.05 0.60 0.60 1.00

1.00 0.15 0.10 0.30 0.30 0.10

0.005 0.00075 0.0005 0.0015 0.0015 0.0005 0.00975

Integrated Microsystems

Table 1 Comparison of Manufacturer Packaging Costs (U.S.$) for a Single-Use Device and for a 200-Dose Clinical Chemistry Kit

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many replicates of test geometry and reagents in a single device, or by running repeat tests in the same set of geometry. Replicating test geometry is limited by the difficulty of assuring device uniformity over large areas, and by requirements for additional interfaces. As with single-use devices, these cannot run quality control specimens to assure full device performance since each section is used only once. The repeat testing scenario requires washing of the device so that residual material from prior analyses does not affect the results of testing. The difficulty of this depends on the analytes to be determined. Table 2 summarizes the allowable proportion, expressed as parts per million, of residual sample material from a prior analysis. The values are calculated by allocating to carryover one half of the allowable imprecision at the low end of the concentration range, assuming contamination by a prior sample at the maximum clinical value of the analyte. The allowable imprecision is taken as the reported low-end imprecision for presently available commercial assays of that analyte. Assay and clinical values are given in Table 2 in the usual reporting units for each analyte. Carryover is expressed as parts per million. The above values show the broad range of cleaning requirements. An assay of bilirubin can tolerate substantial contamination. The most restrictive cases are alpha-fetoprotein (AFP), chorionic gonadotropin, and hepatitis B—analytes with very broad dynamic ranges. In cases where a microsystem is designed to measure more than one analyte, the more restrictive carryover requirement applies. Removal of adsorbed materials from surfaces usually requires mechanical work to rewet the surface and displace the bound material [10] Work may be added by hydraulic flow of rinse fluid, but, in the absence of turbulence, flow rates for pressure driven flow near the walls are close to zero. Driving plug flow by electrokinesis increases shear at the channel wall, but this drive method is dependent on wall properties. When materials are adsorbed they alter the wall charge properties that control electrokinetic flow; plug flow is well controlled only in regions without adsorbed material. This does not mean that electrokinetic flow does not help. The absence of predictable control does not mean the absence of effect; flow rate and mechanical work transfer may even increase

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Table 2

Summary of Allowable Proportion of Redidual Sample Material from a Prior Analysis

Assay Creatine kinase Total bilirubin Creatinine IgG AFP Ferritin hCG hepatitis BsAg

Units

Assay minimum

Clinical maximum

(U/L) (mg/dL) (mg/dL) (mg/dL) (ng/mL) (ng/mL) (mlU/mL) (ng/mL)

5 0.1 0.3 200 3 5 10 5

4,100 30 400 10,000 100,000 5,000 1,000,000 100,000

Allowable imprecision 4 0.15 0.1 10 0.65 0.46 5 0.44

Carryover (ppm) 488 2500 125 500 3 46 3 2

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in regions with adsorbed material. Experience with diluted serum in fused silica capillaries suggests that flow rates are relatively unperturbed. This experience may not be directly transferable to anisotropically etched glass with its rougher surface. Many design features of microfluidic systems make them difficult to clean. In particular, very small concave radii can trap fluid, and abrupt channel size transitions create unswept dead volumes. Long channels of constant cross section avoid unswept dead volumes but have large surface area for their volume. These produce high backpressures that make high-rate flushing difficult. Devices employing anisotropic etching of channels with flat covers have acute angles with small equivalent radii where the cover intersects the trapezoidal channels. These trap residual material if channels are emptied and precipitate buffer salts as they dry. Geometry further contributes to cleaning difficulty because low Reynolds number flow precludes rapid rinse exchange at surfaces. Connections to outside fluids are particularly problematic. Relatively large holes intersecting small channels through the cover create fluid traps. An alternative is edge connection where the channels intersect a free edge of the device [11,12]. This has the added advantage that the interfaces can be mass fabricated. Unlike drilled holes, no individual machining operations are necessary for each connection. Self-sealing polymer channel networks [13] permit a novel approach to cleaning. These may be disassembled between uses and subjected to more rigorous cleaning with the cover removed and the channels exposed. Free access to opened channels removes the laminar flow constraints normally present. Turbulent rinsing of the surface more readily transfers mechanical energy to displace adsorbed materials. Removal of residual prior samples depends on the nature of the channel surface, on the wetted area, on the volume of sample used, and on the nature of the limiting analytes. Fresh hydrophobic surfaces (such as polystyrene) bind many proteins strongly. This may be acceptable provided bound material remains bound and does not influence later measurements. Surfaces may be pretreated with blocking materials to inhibit binding; these blocking agents need to be tested for individual analytes. Clean hydrophilic surfaces have their own problems— glass binds different regions of proteins but clings just as stubbornly.

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This is less of a problem with analytes of narrow dynamic range. Devices intended for ‘‘general chemistry’’ type tests (electrolytes, small metabolites, and substrates) can get by with less rigorous removal of analyte. These are also less likely to adsorb to device surfaces. Cleaning remains a serious roadblock for full menu chemistry. 16.5 SAMPLE VOLUME The history of clinical chemistry has been one of increasingly sensitive methods of analysis employed both to measure lower concentration analytes and to reduce the volume of sample used. Early tests used milliliters of serum for single analytes of millimolar concentration. More recently, sample size has declined to a few microliters (the range of 2–20 µL is common) with sensitivity down to 10⫺14 Molar. Microfluidics offers the capability of measurements with sample sizes from picoliters to nanoliters. Clearly, it is advantageous for the patient to have less blood drawn for analyses, particularly in the case of prolonged illness requiring frequent sampling. Potential reductions of 3–6 orders of magnitude in sample volume for microfluidics are unlikely to be realized in practice. Factors that constrain this include: 1. 2. 3.

4.

16.5.1

Minimum collection volumes associated with sample collection procedures. Dead volumes associated with sample transfer and fluidic interface. The tradeoff between volume and sensitivities—small volumes of low concentration analytes have few molecules. This introduces sampling errors that degrade precision. Adsorption of sample materials onto channel surfaces. This can deplete the sample of analyte, limit detection by nonspecific binding effects, or interfere with electrokinetic fluid control. Sample Collection

Table 3 summarizes various blood collection techniques and the volumes commonly collected. The minimum sample volume is that

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Table 3 Summary of Various Blood Collection Techniques and the Volumes Commonly Collected Collection Minimum sample volume volume (serum (whole blood) or plasma)

Method Venipuncture Capillary (heel or fingerstick) Interstitial fluid

3–10 mL 20–200 µL

0.5 mL 10 µL

2–10 µL

1 µL

Remarks Most commonly used Not commonly automated Not yet available, nonstandard sample

volume of serum or plasma produced from the minimum collection volume of whole blood. A microfluidic device that uses a tiny sample volume provides no benefit to the patient if a large sample is drawn and the bulk is subsequently discarded. Hospitals typically use venipuncture, which is more convenient, has less opportunity for worker exposure, and provides a comfortable excess of sample for sharing between analyzers and for archiving. Venipuncture specimens are drawn into sealed evacuated tubes. The smaller capillary draw requires more labor and more directly exposes the phlebotomist to possibly infectious sample. Interstitial fluid is the extracellular liquid that bathes most tissues. Several companies are preparing minimally invasive devices to sample a small quantity of this fluid. Although this fluid has been shown to reflect plasma levels of small molecules such as glucose with a few minutes’ time constant [14,15], other analytes have not been studied; large protein levels may differ substantially from plasma. 16.5.2

Sample Separation

Conventional analysis requires removal of blood cells to ensure precision, to remove interfering substances, and to prevent contamination of plasma with cell contents or by cell metabolism. Analyses of patient nucleic acids require cell lysis to gain access to the analyte. Since most

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blood cells do not contain a full complement of DNA, lysing all of the blood cells in a sample produces a large mass of debris that presents a clogging hazard. Selecting nucleated cells reduces this problem. The fraction of volume occupied by cells (the hematocrit) varies by up to 20% in normal samples. This sample-to-sample volume variation contributes a like level of uncertainty to assay results; a measured quantity of whole blood contains an uncertain volume of plasma. The size of this uncertainty exceeds the acceptable imprecision for most assays. This may be corrected for either by aliquoting a measured volume of separated plasma, or by determining the hematocrit and adjusting measured concentrations to plasma concentrations. In assays where the volume of sample is not critical, whole blood may be analyzed directly. This is possible where the added uncertainty is not clinically relevant (typically in assays to report the presence or absence of an analyte). It is also possible where the analysis method does not significantly perturb the concentration of analyte in the sample. This requirement is met by equilibrium concentration sensitive assays such as ion-selective electrode determinations of electrolytes without significant sample dilution. It is not met by most conventional clinical assays. The majority of target ions do not directly interact with the ionophores of ion-selective electrodes and optodes. This is possible because the analysis reaction is limited to a small number of analyte receptors at chemical equilibrium. An electrode can produce a detectable signal based on only a small fraction of the high-concentration ions. Larger volumes of sample do not change the measurements because the sensors are already at equilibrium. At low concentration, or with nonequilibrium sensors, a larger fraction of analyte molecules is required to generate a detectable signal. This depletes the sample volume of analyte. A larger volume of sample provides more analyte molecules and changes the measurement results. Such mass-sensitive assays must have cellular elements removed to avoid propagation of the uncertainty in sample volume into assay results. Blood samples are conventionally separated by sedimentation of the cells under centrifugal force. Direct costs associated with the procedure are ⬃ $0.50 per sample. This is not a practical process to implement in a microscale device.

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Filtration or aggregation may conveniently separate small volumes. This is rarely adequate for conventional clinical chemistry because the separated volumes are small, but it fits well with the requirements of microdevices. Blood cell filters have been integrated into microfluidic manifolds [16], but the packed cells have very high nonlinear viscosity and are subject to clotting. These effects conspire to make such filters difficult to clean for subsequent samples. External filters may be cut from bulk material (such as Pall Hemasep) and discarded after use. This is more economical than discarding the entire device. A method unique to the microscale is diffusion-based extraction of analytes away from the cells [17]. This has the benefit that cells remain in dilute suspension and may be relatively easily washed out. Other possibilities are local concentration of cells at nodes of ultrasonic standing waves [18] or diaphoreses [19]. Both provide the possibility of resuspending the cells to rinse them away and are applicable to small active volumes. 16.5.3

Sample Transfer and Loading

Once a sample is collected it must be transferred to the assay device. This usually involves an intermediate container with dead volume that traps and wastes sample. Small-sample droplets cling to the walls of containers; as sample volume decreases, the relative uncertainty in the location of droplets gets larger. If samples are too small, they cannot be located in conventional containers without active targeting. A similar wastage occurs in microfabricated devices. In some cases this is due to mismatch between the size of the channels and fluid load ports. Fluid load ports need to interface with macroscale devices such as hand-guided pipettes or fingertips. Unless carefully designed, these trap much more fluid than actually used in the assay. Some fluid control schemes require extra sample to avoid biasing results. Simple electrokinetic injection methods employed in capillary electrophoresis systems enrich the injected slug for high-mobility species. Microfluidic systems can avoid this problem by designing sample loops [20] that discard the biased material and ‘‘slice off ’’ a slug of sample once the slowest species catch up. The amount of sample discarded is large relative to the volume used; for a 10-fold difference in

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mobility between fastest and slowest species, at least 10 times more sample must flow past the loop than is used. Still more sample must be held in a source reservoir for this injection method, as it only produces an unbiased sample if there is no substantial change in concentration of any species in the source reservoir. The concentration of the highest-mobility species in the sample slug is similar to that in the source reservoir at the end of the injection period. To preserve the concentration of the fastest-moving species to within 1% of the initial sample value requires that ⬍1% of that species leave the source reservoir (this assumes negligible fractional outflow). If the mobility of the fastest analyte exceeds that of the slowest by 10fold, then the source reservoir must contain at least 1000 times the volume of sample actually injected. 16.6 VOLUME SENSITIVITY TRADEOFF As sample volumes decrease, random fluctuations in the distribution of individual molecules decrease the precision of analytical results. Highsensitivity quantitative assays (such as ‘‘third-generation’’ assays for thyroid stimulating hormone) require measurement on the order of 10⫺14 M. The low-end concentration corresponds to six molecules per nanoliter. Assuming every molecule present in a one nanoliter sample were detected, the random distribution of molecules in the specimen contributes a relative standard deviation (expressed as CV) of: √6 ⫽ 41% 6 To contain this Poisson statistics limited contribution to 1% requires 10,000 molecules corresponding to 1.7 µL of sample. Further, given available antibody affinities, realistic reagent concentrations, and acceptable reaction times, only a relatively small fraction of molecules will participate in an analytic reaction. If 10% of the molecules are detected, 100,000 molecules must be in the sample to limit the CV to 1%. This corresponds to a sample size of 17 µL. A channel 50 µm

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wide by 50 µm deep would need to extend more than 6 m to contain this volume. 16.7 ADSORPTION TO DEVICE SURFACES A channel of this configuration has more than six times the surface area of a conventional microtiter well. Most surfaces adsorb components, particularly proteins, from plasma. This surface area is sufficient to significantly deplete low-concentration analytes from a pure solution. Plasma is far from a pure solution and has sufficient amounts of proteins (primarily albumin) to block bulk depletion, but the binding of plasma components to surfaces is not uniform. Some analytes bind more selectively. For example, aminoglycoside antibiotics bind strongly to glass and silica. Without tailored blocking strategies and limited surface exposure, sufficient analyte may be lost to wall adsorption that accurate determinations cannot be made. This becomes more of a problem if the sample is fractionated to remove interfering components. Analyte in protein-depleted fractions may be completely lost to channel surfaces. The blocking of analyte wall adsorption by plasma proteins is not a benign process. If the surface chemical properties of the wall are used to control fluids electrokinetically, then this control can be altered as the walls adsorb plasma components. Since plasma proteins have both hydrophilic and hydrophobic regions, it is difficult to select a material that does not bind proteins. To the extent that the binding is reversible, a coated wall acts as a reservoir of potentially contaminating analyte for later analyses. A further problem related to surface adsorption is nonspecific binding (NSB) in ligand assays. NSB usually dominates the background signal that ultimately limits the sensitivity of low concentration heterogeneous immunoassays. Immunoassays in microfluidic devices need reaction vessels with low surface area per volume to maintain sensitivity. 16.8 SPEED OF ANALYSIS 16.8.1

Requirements

To be useful as a routine diagnostic test, results from an analysis must be complete in a reasonable time. Hospital turnaround times average

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2–4 hr for routine analyses, and one-half hour to 1 hr for stat analyses. The majority of this time is consumed in sample handling; the steps of collection, transport, preparation, data entry, and loading are much longer than the actual analysis. Bringing the analysis closer to the patient can eliminate much of this delay and reduce cost by eliminating handling steps [21]. This is a clear opportunity for microfluidic devices if the promise of more compact fluidics can be realized in more compact instruments. In physicians’ offices and clinics, turnaround time can be hours to days as tests are often sent out to hospital or reference laboratories. Results turnaround during an office visit may save the costs of a second visit. Moving analysis closer to the point of care highlights another potential advantage of integrated microdevices. Specialized instrument operators with time and expertise to perform maintenance, quality control, and instrument troubleshooting are not available at the point of care. Integrated microsystems make it reasonable to replace entire fluidic systems in response to clogs or leaks. This reduces both troubleshooting and maintenance concerns. Such full fluidics replacement is not practical for conventional systems. These point of care applications make sense if the sample handling delays are substantially reduced. The total turnaround time should approximate the present analysis time. The generally means ⬍1 hr; the competitive standard is: ⬍1 min for critical care (blood gases and electrolytes); 5–10 min for routine (metabolites, therapeutic drugs, enzymes, and others above about nanomolar concentration); and 15–30 min for high-sensitivity analytes (concentrations to ⬃ 10⫺14 M). A central lab system needs to achieve or surpass these analysis times. 16.8.2

Capability

Microfluidic devices present some special concerns for speed of analysis. Analysis speed depends on many parameters of a system: 1. Reactions cannot be measured until signals reach minimally detectable levels.

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2. The fraction of analyte available for measurement declines as the analyte concentration decreases. 3. The rate of reaction is limited by exposure of analyte to the detecting reagent; in the low-Reynolds-number environment of microfluidics devices this is dominated by diffusion. Although single fluorophore detection is possible in microdevices, this requires extreme measures difficult to achieve outside of a research environment. Detection of a few hundred fluorophores is more readily attained with practical systems. The slowest common assays are low-concentration ligand-binding reactions. Maximal signal is attained at reaction equilibrium, but this may take many hours to reach. Reactions will asymptotically approach chemical equilibrium so that the majority of the signal is developed in the early stages of the reaction. Care must be taken that the early stages are also the point of greatest sensitivity to reaction conditions. Slight variations in temperature, in mixing history, or in timing can cause large differences in signal. A diminishing fraction of analyte will be bound, and hence detectable, as the concentration decreases. This applies at equilibrium, but also at each point up to equilibrium. The equilibrium distribution of bound versus free analyte is determined by the affinity of the ligands and by the concentrations of ligands and of analyte. This can be controlled by selecting high-affinity ligands and by increasing the ligand concentrations. Although biological ligands have affinities up to 10 14 M ⫺1 and some antibodies can reach 10 12 M ⫺1, most have affinities ⬍10 10 M ⫺1. The maximum concentration of specific antibodies depends on the format of the assay. In solid-phase immunometric (sandwich) assays, the amount of useful capture antibody is limited by the available surface area. Microfabricated devices can produce huge surface areas, but large surfaces increase NSB. NSB is usually the limiting factor for assay sensitivity. High concentrations of reporter antibody, which bind to and identify the captured analyte in the solid phase format, contribute directly to NSB. Directly controllable factors that push reactions to bind large fractions of analyte, and thereby develop large signals, also contribute to large backgrounds that make those signals less distinguishable. This means that there is a practical limit to the usable amount of antibody

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in an immunometric assay. This sensitivity mandated concentration limit can contribute to limitations on the rate of assays. Other rate limitations arise from mass transfer effects. The average separation distance between reactants in part determines reaction rates. At low analyte concentrations, a ligand-coated solid phase depletes the local environment of analyte by binding it. The rate of continuing reaction is limited by the rate that fresh analyte can reach the solid phase. With high concentrations of high-affinity antibodies diffusion limits this rate [22]. This is particularly true when the analyte is a large slowly diffusing molecule, as is usually the case for clinically useful low-concentration analytes. Any presentation of new analyte due to mixing must be of a scale small enough to break up the depleted region. This is difficult in turbulence-free microfluidics devices of low Reynolds number. However, microsystem’s control over details of the geometry affords the opportunity to design short diffusion paths into the device. Alternatively, nonturbulent fine scale mixers [23] may be designed that finely divide the fluid and bring fresh analyte very close to previously depleted reaction zones.

16.9 CONCLUSIONS Clinical chemistry applications present a wide variety of constraints to integrated microfluidic systems. It is relatively easy to apply these new technologies using components derived from other applications so that the composite does not function for clinical assays. Worse, it is possible to produce perfectly functional devices that have no fit to real-world use. Key consequences of the application of clinical chemistry constraints to integrated microdevices are: 1. 2. 3. 4.

Devices Devices Devices Devices

should should should should

accept relatively large sample volumes. be reusable. limit wetted surface area and dead volume. target analysis times of 15 min or less.

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Microfluidic devices must be regarded as components of integrated units, and those units must further be regarded as components of healthcare delivery systems. As the field matures, the integrated microsystems have promise as important elements in the design arsenal.

REFERENCES 1. Erickson, K.A., Wilding, P. Evaluation of a novel point-of-care system, the i-STAT portable clinical analyzer. Clin. Chem. 39(2):283–287; 1993. 2. Connelly, N.R., Magee, M., Kiessling, B. The use of the iSTAT portable analyzer in patients undergoing cardiopulmonary bypass. J. Clin. Monit. 12(4):311–315; 1996. 3. Freaney, R., McShane, A., Keaveny, T.V., McKenna, M., Rabenstein, K., Scheller, F.W., Pfeiffer, D., Urban G., Moser I., Jobst G., Manz A., Verpoorte E., Widmer M.W., Diamond D., Dempsey E, Saez de Viteri F.J., Smyth M Novel instrumentation for real-time monitoring using miniaturized flow systems with integrated biosensors. Ann. Clin. Biochem. 34(Pt 3):291–302; 1997. 4. Verpoorte E., Manz A., Ludi H., Bruno A.E., Maystre F., Krattiger H., Widmer M., van der Schoot H., de Rooij N.F. A silicon flow cell for optical detection in miniaturized total chemical analysis systems. Sensors Actuators B6:66–70; 1992. 5. Bergmeyer H.U., Horder M., Rej R. Approved recommendations (1985) on IFCC methods of measurement of catalytic concentration of enzymes, Part 3. IFCC method for alanine aminotransferase. J. Clin. Chem. Clin. Biochem. 24:481–495; 1986. 6. Horder M., Elser R.C., Gerhardt W., Mathieu M., Sampson E.J. Approved recommendation on IFCC methods for the measurement of catalytic concentration enzymes, Part 7. IFCC method for creatine kinase. Eur. J. Clin. Chem. Biochem. 29:435–456; 1991. 7. Schmidt E., Gerhardt W., Henkel E., Klauke R., Liese W., Wrentz K., Sonntag O., Stein W., Weidemann G. Proposal of standard methods for determination of enzyme catalytic concentration in serum and plasma at 37°C. I. Alkaline phosphatase. Eur. J. Clin. Chem. Clin. Biochem. 30:247–256; 1992. 8. Moss D.W., Henderson A.R. Clinical enzymology. In: Tietz Textbook of Clinical Chemistry, 3d ed. Burtis CA, Ashwood ER, eds. W.B. Saunders, Philadelphia, 1999.

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9. Newark Electronics. Catalog 117. Author, Chicago, 1999. 10. Rosen M.J. Surfactants and Interfacial Phenomena. John Wiley and Sons, New York, 1978. 11. Bell, M.L. Capillary interface and holder. U.S. patent 5,858,194 (1999). 12. Xiang F., Lin Y., Wen J., Matson D.W., Smith R.D. An integrated microfabricated device for dual microdialysis and on-line ESI-ion trap mass spectrometry for analysis of complex biological samples. Anal. Chem. 71:1485–1490; 1999. 13. Delamarche E., Bernard A., Schmid H., Bietsch A., Michel B., Biebuyck H. Microfluidic networks for chemical patterning of substrates: design and application to bioassays J. Am. Chem. Soc. 120:500–508; 1998. 14. Tamada J.A., Bohannon N.J., Potts R.O. Measurement of glucose in diabetic subjects using noninvasive transdermal extraction. Na. Med. 1: 1198–1201; 1995. 15. Smith A., Yang D., Delcher H., Eppstein J., Williams D., Wilkes S. Fluorescein kinetics in interstitial fluid harvested from diabetic skin during fluoroscein angiography: implications for glucose monitoring. Diabetes Technol. Ther. 1:21–27; 1999. 16. Wilding P., Pfahler J., Bau H.H., Zemel J.N., Kricka L.J. Manipulation and flow of biological fluids in straight channels micromachined in silicon. Clin. Chem. 40(1):43–47; 1994. 17. Weigl B.H., Yager P. Microfluidic diffusion-based separation and detection. Science 283:346–347; 1999. 18. Peterson S. Development of an ultrasonic blood cell separator. Proceedings, IEEE 8th Conference Engineering in Medicine and Biology 154– 156; 1986. 19. Yang J., Huang Y., Wang X.B., Becker F.F., Gascoyne P.R. Cell separation on microfabricated electrodes using dielectrophoretic/gravitational field-flow fractionation. Anal. Chem. 71:911–918; 1999. 20. Effenhauser C.S., Manz A., Widmer H.M. Glass chips for high speed capillary electrophoresis separations with submicrometer plate heights. Anal. Chem. 65:2637–2642; 1993. 21. Felder R.A. Robotics and automated workstations for rapid response testing. Am. J. Clin. Pathol. 104(4 suppl 1):S26–S32; 1995. 22. Stenberg M., Stilbert L., Nygren H. External diffusion in solid phase immunoassays. J. Theor. Biol. 120:129–140; 1986. 23. Branebjerg J., Gravesen P., Krog J.P., Nielsen R. Proceedings, Micro Electro Mechanical Systems Workshop, p. 441; 1996.

Index

Acousto-optical deflector (AOD), 17–18 Active microelectronic array system, 223–269 active microelectronic chips and arrays, 224–229 chip/cartridge and electronic hybridization system, 229– 236 electric field-assisted micro/ nanofabrication, 258–266 approach, 260–266 enabling nanotechnology, 258–260 electronic hybridization, aspects of, 236–240 genotyping applications, 240– 252 accuracy and reliability, 245– 248 combined SNP and STR, 248–252 short tandem repeats, 244– 245 single nucleotide polymorphisms, 240–244 introduction, 22–224

[Active microelectronic array system] other diagnostic applications and systems, 252–258 immunoassays and combined assays, 252–254 integrated sample-to-answer systems, 254–258 Advanced nucleic acid analyzer (ANAA), 187, 188, 191, 192, 194–198 Advanced silicon etch process (ASE), 338, 339 Adsorption to device surfaces, 430 Agar-embedded assays, 402 Agarose-based replaceable separation matrix, 173–178 Air displacement systems, 378– 380 Air ducting system, 192 Amplification, 24–30, 255 Anion exchange chromatography, 74–75 (see also Chromatographic methods) Aplysia californica, 152 Applications, 65–68, 69, 77, 83, 147–154, 183–184, 223– 269, 359–362, 395–397 437

438 [Applications] and automation, 83 of biochip technology to HTS, 395–397 and capillary sample introduction, 147–154 chemical sensors, 360 DNA hybridization, 223–269 databases, 69 genotyping, 65–68, 77, 223–269 microanalytical devices, 360–362 pharmacogenomic, 223–269 Assays, 45–50, 149, 287–290, 386–391, 398–402 agar-embedded, 402 bead-staining assays, 401–402 biochemical, 387–389 cell-based, 49, 389–391 development for miniaturized screening, 386–391 enzyme, 45–48, 80–81 immunoassays, 48–49 kinetic, 149 luminescence-based assay, 400– 401 mass-dependent assay, 398–399 PCR, 49–50 proximity-dependent assay, 399–400 single-pass, 287–290 Attachment of DNA, on plastic devices, 306–312 in situ oligonucleotide array, 306–310 postimmobilization processes, 311–312 quality control, 310–311 Automation, 82–83, 169, 179, 313–316, 385–386 versus manual, 179

Index Bacillus anthracis, 198, 199 Basic sample preparation (see also Sample preparation), 71–85 Bead-staining assays, 401–402 Biological warfare (BW), 186 Bioreagents, 374 Blood and body fluid, 79–80, 118–123 Bonding, 342–344 Calculation of back-flow velocity, 213, 215 Capillary action, and surface tension, 41–42 Capillary electrophoresis (CE), 2, 89, 100, 101, 103–105, 108–117, 129–163, 207– 222, 275–277 (see also Interfacing capillaries to microseparation devices; Sheath flow fraction collection) miniaturization of interface, 103–105 multiplexed, 108 sample introduction, 133–135, 141–154 applications, 147–154 channel separation techniques, 141–147 sheath flow fraction collection, 207–222 system design, 131–141 detection methods, 135–141 rectangular channels, 132– 133 Capillary gel electrophoresis (CGE), 97 Capillary isoelectric focusing (cIEF), 210, 214, 217–218

Index Capillary zone electrophoresis (CZE), 129–131, 214–215, 218–219 Cartridge, 229–236, 287–290 (see also Chip/cartridge and electronic hybridization system; Microchips) Catechol, 146 Cell-based assays, 49 Cell release, 149–154 Cell sorting, 20–21 Cetyltrimethylammonium bromide (CTAB), 81 Channel chromatography, 145–147 (see also Chromatographic methods) Channel electrophoresis technique, 134–141 Channel separation techniques, 141–147, 160 2D, 147 Charge-coupled device (CCD), 117, 119, 137–140, 149, 165, 256, 259, 315, 381– 382, 400–401 Chemical libraries (see Miniaturized high-throughput screening of chemical libraries) Chemical sensors, 360 Chip/cartridge and electronic hybridization system, 229– 236, 271–302 cartridge, 287–290 (see also Cartridges; Microchips) Chromatographic methods, 73–75, 145–147 anion exchange, 74–75 channel, 145–147 gas, 2 gel-based, 74–75

439 Clinical chemistry (see Integrated microsystems in clinical chemistry) Combined DNA indexing system (CODIS), 56 Combined SNP and STR genotyping, 248–252 Compounds, 373–374, 408 handling, 408 Computational fluid dynamics (CFD), 363 Conservation of flux, 45 Consumables, 374–375 Control boards, 195 Costs constraints, 419–425 device reuse, 420–425 of microsystems, 419– 420 savings and throughput, 373– 377 beginning the 1536 transition, 376 bioreagents, 374 compounds, 373–374 consumables, 374–375 equipment costs, 375–376 labor, 376 waste disposal, 375 Coulometric efficiency, 145 Customer education, 406–407 Cutting band collection (CBC), 212 Data handling, 391–395 data volume, 393–394 hit follow-up, 395 quality control, 392–393 visualization and data mining, 394–395

440 Deoxyribonucleic acid (DNA) amplification, 255 arrayed on plastic devices, 303– 317 attachment of DNA, 306–312 automation and system integration, 313–316 introduction, 303–304 surface modification, 305–306 target labeling strategies, 312–313 fraction collection, 213–214, 215–216 fragments, separation of, 165–182 genomic templates, 71–79 hybridization, 223–269 plasmid, 76–79 quality for chips and arrays, 83–84 separation, 156–159 (see also Separation) sequencing, and online sample preparation, 87–127 Design, 59–62, 366 of microfabricated reactors, 366 Detection methods, 135–141, 381–384 rectangular channels, 132–133 sample introduction, 133–135 Devices, description of, 36–37, 261, 420–425 (see also Instrumentation and device operation; Interfacing capillaries to microseparation devices; Portable instrumentation for rapid PCR analysis) reuse, 420–425 Dicing and hole drilling, 344–345 Digital imaging, 381–382 Diethylaminoethyl (DEAE), 74

Index Dopamine, 146, 177–178 Double T injector, 130 Duroplastic polymers, 322 E. coli, 257 Elastomeric polymers, 322 Electrochemical array detection, 140–141 Electric-field-mediated separation, 165–182, 258–266 agarose-based replaceable separation matrix, 173–178 approach, 260–266 enabling nanotechnology, 258– 260 introduction, 165–169 membrane-mediated sample loading, 172 migrating fluorophore labeling, 172–173 optical pathway, 170–171 planar gel microchips, 171 separation platform, 169–170 Electrode fabrication, 346–348 Electrokinetics, 38–40, 42 Electronic hybridization, aspects of, 236–240, 271–302 Electro-osmotic flow (EOF), 208 Electropherograms, 97, 115–117, 122 Electrophoretic microdevices, integrated, 1–34 functionality, integrating, 17–32 amplification, 24–30 cell sorting, 20–21 flow control, 21–24 purification, 30–32 introduction, 1–4 microstructures, 4–5 separations, 5–17 multichannel, 11–17

Index Electroplating methods, 328–329 Embossing, 36 Enzyme assays, 45–48, 80–81 Equipment costs, 375–376 Erwina herbicola, 152 Fiber optics, 135–136 1536 transition, 376 Flow control, 21–24 Flow injection analysis (FIA), 142–143 Flow-thru Chip, 271–302 integrated system, 285–299 cartridge, 287–290 fluidics station, 290–299 introduction, 271–274 system, 274–285 advantages, 281–285 concept, 274 materials and properties, 275–278 preparation, 278–281 Fluid distribution, uniform, 292– 294 Fluid movement, 38–42 Fluidics station, 290–299 Fluorophore labeling, 168, 172– 173 Fluorescein di-β-D-galactopyranoside (FDG), 149, 151 Fluorescein mono-β-galactoside (FMG), 149, 151 Fluorescence correlation spectroscopy, 383–384 Fluorescence polarization (FP), 398–399 Fluorescence resonance energy transfer (FRET), 387–388, 399–400 Fluorescent analysis, 235

441 Fluorescent tags, 168, 172–173, 230, 276, 279 (see also Laser-induced-fluorescence) reader, 230 Fraction collection, 213–214, 215–216 Fragments, separation of, 165–168 (see also Separation) Gas chromatography, 2 (see also Chromatographic methods) Gas-phase reactors, 352–353 Gel-based chromatographic methods, 74–75 (see also Chromatographic methods) Gel electrophoresis separations, 11, 167 Gene expression patterns, analysis of, 296–299 Genetic typing, DNA sequencing, and online sample preparation, 87–127 Genomic DNA templates for PCR, 77–79 Genotyping, 7, 68, 77, 87–127, 176, 223–269 accuracy and reliability, 245–248 combined SNP and STR, 248– 252 from blood, 118–123 short tandem repeats, 244–245 single nucleotide polymorphisms, 240–244 Glass capillary arrays, 275–277 Gluing, 342 Green fluorescence protein (GFP), 390 Hand-held nucleic acid analyzer, 200–204

442 Herpes simplex virus (HSV), 11 Hot embossing process, 319–350 Human Genome Project, 87–91, 95 Imaging, in situ, 294 Immunoassays, 48–49, 252–254 and combined assays, 252–254 Inkjet and piezoelectric microdispensing systems, 377–378 Instrumentation and device operation, 62–65, 187–198 (see also Devices, description of; Portable instrumentation for rapid PCR analysis) advanced nucleic acid analyzer (ANAA), 187 control boards, 195 hand-held nucleic acid analyzer, 200–204 miniature analytical thermal cycling, 187 software, 195–198 thermo-optic module, 187–195 Integrated bread board unit (IBBU), 291–292 Integrated electrophoretic microdevices for clinical diagnostics, 1–34 introduction, 1–4 Integrated microfabricated device (IMD), 271 Integrated microsystems in clinical chemistry, 415–435 adsorption to device surfaces, 430 background, 415–417 cost constraints, 419–425 device reuse, 420–425 of microsystems, 419–420 introduction and scope, 415

Index [Integrated microsystems in clinical chemistry] sample volume, 425–429 collection, 425–426 separation, 426–428 transfer and loading, 428–429 special issues, 417–419 (see also Issues for miniaturization) precision, accuracy, and calibration, 417 shelf life, 418–419 temperature control, 417–418 speed of analysis, 430–433 capability, 431–433 requirements, 430–431 volume sensitivity tradeoff, 429–430 Integrated sample-to-answer systems, 254–258 Integrating functionality, 17–32 Interfacing capillaries to microseparation devices, 129–163 (see also Devices; Instrumentation and device operation) capillary sample introduction, 141–154 applications, 147–154 channel separation techniques, 141–147 channel separations for analysis of STR, 160 introduction, 129–131 separations of DNA restriction fragments, 156–159 system design, 131–141 detection methods, 135–141 rectangular channels, 132–133 sample introduction, 133–135 ultrathin slab gels, 155–156

Index Issues for miniaturization, 403– 408, 417–419 compound handling, 408 customer education, 406–407 obsolescence cycles, 407–408 precision, accuracy, and calibration, 417 productivity, 405–406 shelf life, 418–419 size, 403–404 temperature control, 417–418 timing, 404–405 Kinetic assays, 149 Lab-on-a-Chip, 3, 35–54, 223, 254, 294, 319, 363 assays, 45–50 cell-based, 49 enzyme, 45–48 immunoassays, 48–49 PCR, 49–50 devices, description of, 36–37 introduction, 35–36 microfluidics, 38–45 chip length scales, 44–45 conservation of flux, 45 fluid movement, 38–42 mixing, 42–44 versus array technologies, 51–52 sample and compound accession, 51 Labor, 376 Lamination, 342 Laser ablation, 36, 343 Laser-induced-fluorescence (LIF), 62, 135, 136, 138 (see also Fluorescent tags) Laser welding, 344

443 Light-emitting diode (LED) structures, 262–266 Liquid-phase reactors, 353–356 Low-ionic-strength solutions, 96 Low positive voltage, 64 Luminescence-based assay, 400– 401 Macroporous silicon, 277–278 Mammalian tissue, 80 Mask, 4 Mass-dependent assay, 398–399 Membrane-mediated sample loading, 172 Micellar electrokinetic channel chromatography (MEChC), 145–147 Microanalytical devices, 360–362 Microchips, 3, 5–11, 189 (see also Cartridges; Chips) industry, 3 separations, 5–11 Microdevice design, 59–62 Microelectromechanical systems (MEMS), 189, 223 Microfabricated electrophoretic devices, 55–70 genotyping applications, 65–68 instrumentation and device operation, 62–65 introduction, 55–56 microdevice design, 59–62 microfabrication, 56–59 outlook, 69 Microfabricated reactor technology, 331–370 analytical aspects, 359–363 chemical sensors, 360 microanalytical devices, 360– 362

444 [Microfabricated reactor technology] design, 366 introduction, 351–352 reactors, 352–359 gas-phase, 352–353 liquid-phase, 353–356 packed-bed, 356–357 unit operations, 357–359 simulation of systems, 363–366 techniques, 362–363 Microfluidic technologies (see also Lab-on-a-Chip), 35–54, 319–350 chip length scales, 44–45 conservation of flux, 45 fluid movement, 38–42 mixing, 42–44 by polymer hot embossing, 319–350 additional technologies, 342– 346 examples, 336–341 fabrication, 327–330 introduction, 319–321 materials, 312–323 outlook, 346–348 process, 330–336 replication technologies, 323– 327 versus array technologies, 51– 52 Micromachining methods, 327– 328 Microseparation devices (see Interfacing capillaries to microseparation devices) Microscale devices, 261 Microstructures, in glass microchips, 4–5

Index Migrating fluorophore labeling, 172–173 (see also Fluorescent tags) Miniature analytical thermal cycling instrument (MATCI), 187, 188, 191, 192, 193, 195, 197 Miniaturization of CE interface, 103–105 Miniaturization of the purification column, 107–108 Miniaturization of Sanger reaction, 105–107 Miniaturized high-throughput screening of chemical libraries, 371–414 application of biochip technology to HTS, 395–397 assay development for miniaturized screening, 386–391 biochemical, 387–389 cell-based, 389–391 cost savings and throughput, 373–377 beginning the 1536 transition, 376 bioreagents, 374 compounds, 373–374 consumables, 374–375 equipment costs, 375–376 labor, 376 waste disposal, 375 data handling, 391–395 data volume, 393–394 hit follow-up, 395 quality control, 392–393 visualization and data mining, 394–395 examples, 397–403 agar-embedded assays, 402

Index [Miniaturized high-throughput screening] bead-staining assays, 401– 402 luminescence-based assay, 400–401 mass-dependent assay, 398– 399 proximity-dependent assay, 399–400 introduction, 371–373 issues for miniaturization, 403– 408 compound handling, 408 customer education, 406–407 obsolescence cycles, 407–408 productivity, 405–406 size, 403–404 timing, 404–405 new tools, 377–386 automation, 385–386 detection systems, 381–384 pipetting systems, 377–381 plate formats, 384–385 Miniaturized standard readers, 381 Molding tools, fabrication of, 327–330 electroplating methods, 328–329 micromachining methods, 327– 328 silicon, 329–330 Multichannel genotyping, 68 Multichannel separations, on microchips, 11–17 Multiplexed eight-channel STR, 60–61 Multiplexed integrated online sample preparation, 87–127 (see also Sample preparation)

445 Multiplexed online PCR analysis, 118–123 Multiplexed online reaction and electrophoresis, 108–117 Nanochip cartridge, 229–236 Nanofabrication applications, 223– 269 Nanoscale components, 261 Nanotechnology, 258–260 Ninety-six-channel radial array microchip, 15–16 Nonionic detergent, 81 Nonspecific binding (NSB), 430, 432 Obsolescence cycles, 407–408 Online PCR analysis, 97–103 Online sample preparation, 87– 127 (see also Sample preparation) Online Sanger reaction and DNA sequencing, 91–97 Optical pathway, 170–171 Optics, focusing and collecting, 192–195 Organic extraction, 72–73 Packed-bed reactors, 356–357 Peak activated collection (PAC), 212 Peltier heating/cooling technology, 28–29 Pharmacogenomic applications, 223–269 Photodiode array detection, 135– 136 Photomultiplier tubes (PMT), 62 Physical properties, of polymers, 321–323, 324

446 Piezoelectric and inkjet microdispensing systems, 377–378 Pin transfer technologies, 380– 381 Pipetting systems, 377–381 Planar gel microchips, 165–182 Plant chloroplast, 259 Plant material, 80–81 Plasmid DNA, 76–77 Plastic devices, DNA arrayed on, 303–317 attachment of DNA, 306–312 automation and system integration, 313–316 introduction, 303–304 as substrates, 304–305 surface modification, 305–306 target labeling strategies, 312– 313 Plate formats, 384–385 Polybutylenterephthalate (PBT), 322, 324 Polycarbonate (PC), 322, 324, 325, 333, 334 Polyethylene (PE), 323 Poly(ethylene oxide) (PEO), 91 Polymerase chain reactions (PCR), 3, 7–12, 24–32, 49–50, 77–79, 81–82, 89, 118– 123, 165, 183–206, 210 (see also Portable instrumentation for rapid PCR analysis) amplification, 184 fragments, 81–82 genomic DNA templates, 77–79 impurities and inhibitory effects, 78 rapid analysis, 183–206 submicroliter, 50

Index Polymer hot embossing, and microfluidic devices, 319–350 (see also Hot embossing) additional technologies, 342–346 examples, 336–341 fabrication, 327–330 introduction, 319–321 materials, 312–323 outlook, 346–348 process, 330–336 replication technologies, 323–327 Polymer materials, 321–323 properties, 321–323 Polymethylmethacrylate (PMMA) substrates, 36, 322–326, 331, 333, 334, 336–341, 345 Polyvinylpolypyrrolidone (PVPP), 81 Portable instrumentation for rapid PCR analysis, 183–206 instrumentation, 187–198 control boards, 195 software, 195–198 thermo-optic module, 187–195 introduction, 183–187 next-generation instrument, 200–204 performance, 198–200 Postimmobilization processes, 311–312 Precision, accuracy, and calibration, 417 Pressure, 41, 42, 109 Probe capture, 234 cleavage, 186 distribution, 284–285 hybridization, 186 loader, 230, 231 Productivity, 405–406

Index Properties, of polymers, 321–323, 324 Proteinase K, 78, 79 Protein fraction collection from cIEF separations, 214, 217–218 Proximity-dependent assay, 399– 400 Purification and extraction, 30–32 Purification column, miniaturization of, 107–108 Purity check, 216 Quality control, 83–84, 310–311, 392–393 Rapid genotyping, 176 Rapid PCR analysis, 183–206, 361 (see also Portable instrumentation for rapid PCR analysis) Reaction chamber, 189 Reaction rates, enhanced, 283– 284 Reactors, 352–359 (see also Microfabricated reactor technology) design of, 366 gas-phase, 352–353 liquid-phase, 353–356 microfabricated, 352–359 packed-bed, 356–357 unit operations, 357–359 Rectangular channels, 132–133 Real-time PCR, 200 Replication technologies, 323– 327 Reversible syringe pumps, 109, 120 Ribonucleic acid (RNA), 75

447 Sample and compound accession, 51 Sample preparation, 71–85, 87– 127, 254–258 (see also Interfacing capillaries to microseparation devices) anion exchange chromatography, 74–75 automation, 82–83 blood and body fluid, 79–80 chromatographic methods, 73– 74 current challenges, 71–72 DNA quality for chips and arrays, 83–84 genomic DNA templates for PCR, 77–79 integrated sample-to-answer systems, 254–258 mammalian tissue, 80 miniaturization of CE interface, 103–105 miniaturization of the purification column, 107–108 miniaturization of Sanger reaction, 105–107 multiplexed integrated online, 87–127 multiplexed online PCR analysis, 118–123 multiplexed online reaction and electrophoresis, 108–117 online Sanger reaction and DNA sequencing, 91–97 online PCR analysis, 97–103 organic extraction, 72–73 PCR fragments, 81–82 plant material, 80–81 plasmid DNA, 76–77 silica-based methods, 75–76

448 Sample volume, 425–429 collection, 425–426 separation, 426–428 transfer and loading, 428–429 Sanger reaction and DNA sequencing, 91–97 miniaturization of, 105–107, 112, 113, 123 Scanning confocal imagers, 382– 384 Self-sealing polymer channel networks, 424 Separation of DNA restriction fragments, 156–159 electric-field-mediated, 165–182 on a microchip, 5–11 multichannel, 11–17 in nanometer structures, 145 platform, 169–170 sample, 426–428 Sequencing by hybridization (SBH), 303 Serum cortisol immunoassay, 6 Sheath flow fraction collection in capillary electrophoresis, 207–222 introduction, 207–213 materials, 213–215 calculation of back-flow velocity, 213 DNA fraction collection, 213–214 protein fraction collection from cIEF separations, 214 sugar fraction collection from CZE separations, 214–215 methods, 215–219 calculation of back-flow velocity, 215

Index [Sheath flow fraction collection] DNA fraction collection from polymer-filled columns, 215–216 protein fraction collection from cIEF separations, 217–218 sugar fraction collection from CZE separations, 218–219 Shelf life, 418–419 Short tandem repeat analysis, 55– 70, 160, 166, 240–245, 310 genotyping applications, 65–68, 240–244 instrumentation and device operation, 62–65 (see also Devices; Instrumentation) introduction, 55–56 microdevice design, 59–62 microfabrication, 56–59 outlook, 69 Sidewall angles, 335 Sidewall roughness, 335 Silica-based methods, 75–76 Silicon micromachining, 329–330 Single-channel microdevice, 61, 63 Single nucleotide polymorphisms (SNP), 240–244 Single-pass assays, 287–290 Sodium dodecyl sulphate (SDS), 23, 72–73 Software, 195–198 Speed of analysis, 430–433 capability, 431–433 requirements, 430–431 Spot density, 284–285 Sugar fraction collection from CZE separations, 214–215, 218–219

Index Surface modification, 305–306 Surface tension, and capillary action, 41–42 System for active microelectronic array, 223–269 active microelectronic chips and arrays, 224–229 chip/cartridge and electronic hybridization system, 229– 236 (see also Chip/cartridge and electronic hybridization) electric field-assisted micro/nanofabrication, 258–266 approach, 260–266 enabling nanotechnology, 258–260 electronic hybridization, aspects of, 236–240, 271–302 genotyping applications, 240– 252 (see also Genotyping) accuracy and reliability, 245– 248 combined SNP and STR, 248–252 short tandem repeats, 244– 245 single nucleotide polymorphisms, 240–244 introduction, 22–224 other diagnostic applications and systems, 252–258 immunoassays and combined assays, 252–254 integrated sample-to-answer systems, 254–258

449 System design, for channel electrophoresis, 131–141 (see also Design) detection methods, 135–141 rectangular channels, 132–133 sample introduction, 133–135 Tamoxifen, 297, 298 Target labeling strategies, 312– 313 T assembly, 110, 118, 119 Temperature control, 295–296, 417–418 Temperature sensor, 191 Thermal load, 28–29 Thermal mass constraints, 29 Thermo-optic module, 187–195 Thermoplastic polymers, 322 Thin film lamination, 36 384-channel pin tool, 379 Ultranarrow channels, 141 Ultrasonic welding, 344 Ultrathin slab gels, 155–156 Unit operations, of reactors, 357– 359 Voltage, 64 Volume sensitivity tradeoff, 429– 430 Waste disposal, 375 Water gas shift (WGS) reaction, 356–357 Yersinia pestis, 198, 200