Sample pretreatment on microfabricated devices

Talanta 56 (2002) 233– 266 www.elsevier.com/locate/talanta

REVIEW

Sample pretreatment on microfabricated devices Jan Lichtenberg, Nico F. de Rooij, Elisabeth Verpoorte * SAMLAB, Institute of Microtechnology, Uni6ersity of Neuchaˆtel, CH-2007 Neuchaˆtel, Switzerland Received 24 September 2001; received in revised form 12 October 2001; accepted 15 October 2001

Abstract The integration of sample pretreatment into microfluidic devices represents one of the remaining hurdles towards achieving true miniaturized total analysis systems (mTAS). The challenge is made more complex by the enormous variation in samples to be analyzed. Moreover, the pretreatment technique has to be compatible with the analysis device to which it is coupled in terms of time, reagent and power consumption, as well as sample volume. This review provides a thorough overview of the developments in this field to date. © 2002 Elsevier Science B.V. All rights reserved. Keywords: Sample pretreatment; Microfabricated devices; Miniaturized total analysis systems; Sample preparation; Purification; Filtering; Preconcentration; PCR; Dialysis; Labeling

1. Introduction Analytical chemistry is a broad field, with many different methods of measurement based on assessing a variety of characteristics which can differentiate one species from another. Fundamentally, though, all analytical procedures follow a certain set of processes in a well-defined sequence. This starts with a sampling step, in which a (hopefully) representative sample is obtained from the system about which chemical information is desired. The sample then usually undergoes some kind of sample preparation or pretreatment step before being submitted to the actual analysis. This step may involve extracting the sample from its * Corresponding author. Tel.: +41-32-7205-442. E-mail address: [email protected] (E. Verpoorte).

matrix, removing large matrix components from the sample, masking or removing species which could interfere with the measurement, derivatization to make the sample detectable, or a sample preconcentration step. Once the sample’s condition is suitable for the analysis method chosen, the sample is subjected to analysis, at which time the presence of the analyte of interest is ascertained and its concentration often quantified. This chemical information is then converted to an electronic signal during the data acquisition step. The goal of analytical chemists over the years has been to make their jobs easier, through au-tomation of as many of the steps described above as possible. The total chemical analysis system (TAS) [1], and more recently, the miniaturized total chemical analysis system (mTAS) [2], have been concepts put forth to address this issue. The TAS proposed

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the full incorporation of analytical procedures into flowing systems, with carrier streams transporting sample from one manipulation to the next. The mTAS is a smaller, faster version of this, with analyses taking place in flow systems having microliter and even sub-microliter volumes, to achieve analysis times on the order of seconds rather than many minutes. The devices used are small and monolithic in nature, with cross-sectional dimensions on the order of tens of micrometers. Since the first demonstrations of fast capillary electrophoresis (CE) separations on chip in the early 1990s [3– 5], researchers have developed a basic set of microfluidic tools which can be successfully applied to define and move small volumes of sample, as well as mix them or dilute them, as the case may be. In other words, the basic liquid manipulations we associate with an analysis (aliquoting, adding a reagent, mixing, etc.) can, for the most part, be accomplished on a chip. The most common fluid pumping mechanism implemented to date has been electro-osmosis, using electric fields to generate a bulk flow in conducting liquids [6]. The first examples of on-chip mixing and dilution came soon after the initial demonstrations of CE, accompanied by other papers describing onchip derivatization of samples for easier detection by fluorescence. The trend since then has been towards increasingly complex microfluidic networks, for enhanced sample handling capacity through integration of several interconnecting elements into the same device. An excellent analysis of this for electrokinetically driven devices may be found in Ref. [7]. However, the day of the true mTAS, systems capable of dealing with raw samples no matter what their composition, is still some way off. One of the main challenges that remains for mTAS researchers is chip-based sample pretreatment. For one thing, it is not always trivial to move nanoliter or picoliter samples around in a branched microfluidic device, an essentially open system in which solutions of different compositions must be localized to different areas of the chip. Hence, integration of different sample pretreatment steps is not just a matter of joining microchannels on a device. It is still more practi-

cal to perform some of the sample pretreatment steps off-chip in many cases. This situation is changing rapidly, though, as more insight is gained into the behavior of fluids in microchannels. The other issue standing in the way of complete integration of sample analysis onto a chip is the highly complex nature of many of the samples that need to be analyzed. To date, the samples dealt with in microdevices have been quite pristine for the most part, having undergone some form of manual sample preparation primarily to avoid clogging of the microscopic features in the chip. Though it may seem contradictory, the ability of microchannel-based devices to handle samples containing particles or strongly adsorbing molecular species will be essential to the full realization of the mTAS concept. An examination of the literature reveals a variety of examples for chip-based sample pretreatment, and it becomes clear that there is no one universal approach to this challenge. This is, of course, not surprising, given that sample pretreatment procedures have always had to be individually tailored to the type of sample under consideration and the analytical method chosen. Generally speaking, though, chip-based methods can be placed in one of four major categories, analogous to their conventional counterparts. These categories are outlined in Fig. 1 and described below, in more or less the chronological order of procedures any given sample would undergo, starting from the sampling step.

1.1. Separation of sample from sample matrix In most cases, real-world samples are not suitable for direct analysis for a number of reasons. Often, samples contain large organic or inorganic particles, which must be removed to prevent disruption of fluid handling by fouling or blockage of the analytical system. In bioanalytical applications, the sample is often contained within a cell, and must be removed by means of some cell membrane rupture technique. In all these cases, sample is being separated from a complex background matrix in an initial purification step.

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1.2. Sample preconcentration

1.4. Biochemical sample pretreatment

If the sample analyte is only available in trace amounts, detection of a signal over the background noise becomes difficult if not impossible. Preconcentration of sample prior to analysis can alleviate the sensitivity demands placed on the detector, by squeezing available analyte molecules into a smaller volume for enhanced detection. On-chip, these techniques are primarily used in conjunction with separation methods.

Biochemists have a large set of additional reaction tools at hand to transform large molecules into more easily handled entities. Amplification of DNA using the polymerase chain reaction (PCR) and cleavage by restriction enzymes are two prominent techniques widely used in genomics, whereas enzymatic digestion of proteins is established for protein analysis. There has been a surge in the development of chip-based methods for DNA and protein analysis, commensurate with the increased demand for information in genomics and proteomics.

1.3. Deri6atization Derivatization involves the chemical transformation of the analyte to render it detectable by the detection system used. In the biochemical domain, specific or non-specific labeling of biomolecules with fluorescent labels is a common technique. A multitude of flow injection analysis methods are based on classical wet chemical methods, to convert the species of interest into a colored compound for detection by optical means. A significant number of examples for sample derivatization exist in chip-based analysis, to facilitate small-volume detection particularly by fluorescence.

This article seeks to review important developments published to date in the area of chip-based sample pretreatment, following the outline depicted in Fig. 1. The discussion will focus primarily on examples of liquid sample pretreatment, as these predominate in the mTAS field. The categorization of these examples will follow that laid out above, but in a somewhat different order, determined by the degree of complexity of the fluid or sample handling involved. Thus, chip-based derivatization will be presented first, since this is generally accomplished by relatively simple

Fig. 1. Classification of important sample pretreatment techniques.

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reagent additions in branched microchannel devices. Sample preconcentration techniques on chips are then covered, followed by more complex devices for biochemical amplification, sample filtration, and the like. To start, a few examples for preconcentration of gaseous analytes on chip will be described.

motic pumping. This requires conductive liquids containing ions, and works best in aqueous systems.

3.1. Labeling and complexation

3. Sample preparation for liquid samples

Fluorescence detection continues to be the method of choice for chip-based analysis, as it has superior sensitivity and is ideally suited to ultrasmall volume (pl) analysis and/or detection of single molecules. However, most species are not intrinsically fluorescent, and so must be covalently labeled with a fluorescent tag. Generally, these tagging reactions are carried out manually before the separation is done, which leads to additional preparative tasks and the requirement for comparatively large sample volumes. In addition, the total analysis times are drastically increased if labeling times are taken into account. For instance, while the first reported separations of fluorescein isothiocyanate (FITC)-labeled amino acids took only a few seconds, the labeling reaction beforehand took several hours [9]. There do exist fluorescent labeling reactions for aminated species that are kinetically very fast, however, opening up the possibility of on-line tagging directly before or after analysis by separation. This type of application is ideal for a chip-based approach, since low dead-volume connections between microchannels minimize band broadening due to mixing with the reagent. Alternatives to covalent fluorescent labeling are available for post-column (post-separation) tagging of species. These include non-covalent fluorescent labeling, complexation and chemiluminescence. Flow injection analysis-type methods have been integrated into microfluidic devices having larger volumes, on the order of microliters, for conversion of inorganic, ionic species to colored molecules detectable by absorbance, fluorescence or other optical methods. In this case, no separations are carried out.

At present, the majority of microanalytical systems focus on liquid samples which are primarily aqueous. This is in part due to the application needs in biochemistry. It is probably also a result of the fact that the most convenient method for managing liquids in nanoliter microchannels is electro-os-

3.1.1. Post-column reactors for labeling The first example for on-chip labeling reactions was a post-column fluorescent tagging of amino acids using o-phthaldialdehyde (OPA) [10]. This particular label was chosen for its rapid reaction kinetics. The planar glass device consisted of a

2. Sample preparation for gaseous samples Only a few explicit examples of miniaturized sample preparation techniques for gaseous samples have been published, although generally many micromachined filters would also be suitable for removing particles from gas streams. These types of filters will be presented later. An interesting example of preconcentration of a gaseous sample prior to on-chip chromatographic analysis has been published by Sandia National Labs [8]. The device consists of a flow-through chamber with an integrated micro-hotplate covered by a mesoporous adsorbant sol– gel layer. At room temperature, this layer collects the analyte of interest very efficiently, while interferents are not well adsorbed, leading to a selective preconcentration. The gaseous species focused on in this work was dimethyl methyl phosphonate (DMMP), a substance used as nerve gas simulant. Once the sample collection is finished, a short (100– 200 ms) heating pulse of 200 °C applied to the micro-hotplate rapidly releases the DMMP, creating a sharp input plug to the gas chromatograph. A 5× 8 mm preconcentrator achieves signal gains of up to 300-fold for a 90 s collection time. Due to the small area and low thermal mass of the micro-hotplate, heating rates of 104 °C s − 1 can be achieved, while maintaining a low power consumption of 100 mW.

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Fig. 2. (a) Layout of a microchip with integrated post-column reactor for fluorescent labeling of separated amino acids as proposed in [10]; b) a design optimized for rapid mixing and labeling reaction leading to improved separation efficiency [12] (reprinted with permission from [10] and [12]. Copyright 1994 and 1996 American Chemical Society).

typical microchip CE structure with an additional side channel intersecting the separation channel at a right angle 6 mm after the injection cross (Fig. 2a). Via this side channel, the labeling reagent was brought into contact with the separated analyte bands. Reaction took place along the way downstream, followed by fluorescence detection at 351.1 nm using an argon-ion laser. Band broadening due to reagent addition at the entrance of the reaction column was studied by

injecting rhodamine B plugs into the system and comparing peak shapes before and after the mixing point. For electrical field strengths above 500 V cm − 1, the measured plate height, H, increased by a factor of 2. However, the separation of post-column-labeled amino acids was much less efficient, H being 3–23 times larger than the values for rhodamine B. This additional band broadening was attributed to the different migration speeds at which already-labeled and non-la-

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beled analytes migrated in the channel. Although the kinetics of the labeling reaction were relatively fast (the reaction half-time is 4 s), analyte bands spread in the reaction column. Harrison et al. worked on post-column reactors using the same sample/label combination as Jacobson et al. Initially, they encountered comparably low separation efficiencies using a design with the separation and reaction channels laid out in a straight line [11]. However, a closer analysis of the band broadening source led to an improved design and operation conditions, which allowed separations having theoretical plate numbers, N, of over 83000 [12]. The authors proposed two reasons for band broadening: (1) as in Ref. [10], the analyte band spread due to different mobilities of labeled and unlabeled species during the reaction; (2) differences in electro-osmotic velocity, which are due to mismatched pH or ionic strength in running CE buffer and labeling streams, caused Poiseuille flow profiles and distorted the bands further [13]. In an early configuration similar to that in Ref. [10] (Fig. 2a), the labeling reagent stream was directed into the reaction channel in a perpendicular, head-on manner. This confined the label to a narrow zone at one side of the channel and increased the time necessary for diffusive mixing [11]. In a later design (Fig. 2b), the label and separation channels intersected in a Y configuration, assuring fast diffusive mixing of sample and reagent flows. The second important issue, namely the mismatch in electro-osmotic mobilities of the different liquids, could be verified by changing the pH of the labeling reagent. With a running buffer pH of 9.70 and a labeling reagent pH of 9.65, an N of up to 83000 could be achieved. Adjusting the labeling reagent pH to 8.99 caused band broadening leading to a reduction of N to 20000.

3.1.2. Pre-column reactors for labeling OPA was also the labeling reagent of choice used for precolumn labeling of amino acids on a microchip [14]. In this case, the amino acid sample was mixed with the labeling reagent in a 1 nl reaction chamber before it was injected into the 15.4 mm separation column by time-based, gated injection with 1.8% RSD in peak area (Fig. 3a).

The series combination of reaction chamber and separation channel created a direct dependence of the reaction time for each particular analyte on its electrophoretic mobility. For instance, for a separation field strength, Esep, of 1.8 kV cm − 1, the reaction time for arginine was 4.1 s, whereas the lower mobility glycine had a longer reaction time of 8.9 s. This in turn leads to different labeling efficiencies for each analyte. Consequently, it could be shown that the mass detection limits depended strongly on reaction time and separation field strength, with minimum values of 0.55 fmol for arginine and 0.83 fmol for glycine (Fig. 3b). However, the plate height was still comparatively high (ca. 10–30 mm, depending on Esep), a phenomenon attributed to both the gated injection with its long sample plugs and the roughness of the channel walls due to fabrication limitations. Fluorescent labeling of amino acids prior to off-chip HPLC separation has been proposed using NBD-F (4-fluoro-7-nitrobenzofurazan) as fluorophore [15]. The labeling reaction requires the sample-NBD-F mixture to be heated to 60 °C for 2 min, which is achieved in a 100 mm long, 50 ml volume reaction channel in a silicon–glass sandwich structure. Integrated Cr–Pt resistors heated the chip at a rate of 2 °C s − 1 up to 90 °C, consuming 2 W. The temperature could also be monitored by integrated resistive sensors.

3.1.3. Post-column reactors for complexation Fluorescent tagging is not the only technique to make ions of interest visible for optical detection methods. Kutter et al. presented a microdevice for metal cation analysis combining on-chip complexation and sample stacking, a preconcentration technique which will be dealt with later in this review [16]. Using the same chip structure as in Ref. [10], calcium and magnesium were labeled after separation using 8-hydroxyquinoline-5-sulfonic acid (HQS) to form a fluorescent complex [17]. Fluorescence is maximal at excitation around 390 nm and pH 8, conditions that could be met by using an argon-ion laser and a borate buffer system. A four-fold decrease in signal intensity between off-line and on-line complexation could be observed, which was easily compensated for by field-amplified stacking and injection.

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Liu et al. [18] have combined on-chip CE with post-column complexation using a fluorogenic dye, NanoOrange, to detect proteins by laser-in-

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duced fluorescence. An advantage of NanoOrange is the short reaction half-time of only 110 ms for non-covalent binding to the hydrophobic regions

Fig. 3. (a) The CE microchip with pre-column labeling has an integrated, 1-nl reaction chamber in front of the injection element [14]. (b) The mass detection limits for arginine ( ) and glycine ( ) decrease with increased residence time in the reaction chamber as more labeled product is generated. Residence time in turn depends on the electro-osmotic flow velocity and therefore on the applied electric field (reprinted with permission from [14]. Copyright 1994 American Chemical Society).

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of proteins (compared with 12 s for OPA). The labeling rate is therefore within the range for a diffusion-limited reaction, leading to plate heights of 10 –20 mm.

3.1.4. Post-column reactors for chemiluminescence A different tagging technique for biomolecules integrated on-chip is chemiluminescence detection, based on the horseradish peroxidase (HRP)catalyzed reaction of luminol with peroxide in a post-column scheme [19]. The device used for this experiment is the same as described in Ref. [12], with the added feature of a sputter-deposited aluminum mirror on the back side of the detection zone of some devices for increased concentration sensitivity. Luminol (5 mM) was added to the separation buffer itself, while 1.5% peroxide was mixed with the analyte bands after separation at the Y-shaped intersection prior to detection. The initial optimization of the detector setup showed a 1.6-fold improvement in detection limits for fluorescein-labeled HRP from 171 nM down to 105 nM when using the mirror. Depending on the injection technique used, the separation efficiency was determined to be 1000– 5800 theoretical plates, a value in general sufficient for the immunoassay application of the device. A direct immunoassay of mouse immunoglobulin G (IgG) using an F(ab%)2 fragment of the HRP-conjugate of goat anti-mouse IgG was demonstrated for a concentration range from 0 to 60 mg ml − 1, by injecting the sample after off-chip immunological reaction. 3.1.5. Miniaturized flow injection analysis (vFIA) A number of publications have described the integration of classical wet chemical methods into micromachined fluidic systems. One early example focused on the analysis of phosphate using the molybdenum blue method to form a blue-colored phosphate complex which could easily be detected by absorbance [20,21]. The microfluidic system was three-dimensional in nature, consisting of a number of silicon chips (23×23 mm) stacked on top of one another. Passage of liquid between layers was assured by the regular arrangement of through-holes around the chip perimeter. The overall volume of the system was relatively large,

at 90 ml or so. This example was interesting in that it was the first to use silicon-based micropumps for controlling liquids within the valveless system. A more recent example of mFIA is based on three-layer, glass– silicon–glass devices, containing an integrated optical cuvette etched through the silicon chip [22]. The device was developed for analysis of ammonia in waste and drinking waters, using the Berthelot reaction to convert ammonia through reaction with phenol to a blue indophenol dye. In this case, syringe pumps were used to control flows in the chip, which had an overall volume between 2 and 10 ml. An allglass version of the three-layer chips for this application has also been reported [23]. Electrokinetically driven mFIA systems based on glass have also been used for phosphate analysis by the molybdenum blue method [24], as well as analysis of other inorganic analytes [25].

3.2. Sample preconcentration As pointed out above, fluorescence detection is still the predominant mode used in conjunction with microchips, due to its high sensitivity. However, most species are not intrinsically fluorescent, and many of these are not easily labeled. Hence, ongoing research efforts looking at the on-chip implementation of UV–visible absorbance detection and electrochemical methods, among others, will certainly expand the range of analyses possible. However, none of these methods measure up to fluorescence in terms of achievable detection limits. Many applications will therefore require some form of sample preconcentration prior to analysis to ensure that trace amounts of analytes can be reproducibly quantified. Physiological samples present a particular challenge, since analytes are often present at sub-micromolar levels. The analysis of hormones in serum by liquid chromatography is one such example, where concentrations of analyte range from nanomolar to picomolar, but fluorescent labeling is generally not possible. Though UV absorbance may be used, it exhibits detection limits of at best 10 − 7 M. This performance is severely compromised on-chip and in fused silica capillaries by very short optical pathlengths.

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Table 1 Efficiencies of various on-chip preconcentration concepts

low buffer concentrations, the relative conductivity, k, is defined as:

Technique

Concentration enhancement factor

References

k= cr/cs = |r/|s = Es/Er

Field-amplified sample stacking Field-amplified injection Stacking of neutral analytes Solid phase extraction Surface affinity reactions Porous membrane structures

10–100

[31,32]

10–20

[34,16]

20

[40]

80–500

[46,48,50]

Up to 30

[59]

Up to 100

[53,54]

To counteract lower detection sensitivity, a number of solution-based, on-line preconcentration techniques have been developed for electrokinetic separation. On the microscale level, research work has been undertaken since the mid-1990s to implement some of these methods into planar analysis devices. Other applications involve solidphase extraction of species onto a surface, either selectively or non-selectively, for further analysis by separation or other means. Table 1 summarizes the expected performances of the different techniques, which are discussed in more detail below.

3.2.1. Field amplification stacking techniques Field amplification stacking (FAS) in conventional capillary systems was first mentioned by Mikkers et al. [26] and was intensively studied by Chien, Burgi and coworkers [27– 30]. The technique has been used on microchips for stacking of analytes in 400 mm long, volume-defined sample plugs [31] and for stacking full column’s worth of sample [31,32]. Preconcentration is achieved by generating a high electrical field within an injected sample plug, that rapidly drives and stacks sample ions at the ends of that plug. This field amplification is created when the sample is dissolved in a buffer that has a much lower conductivity than the surrounding running buffer used for separation. Fig. 4 illustrates the mechanism schematically. Assuming

(1)

where cr and cs are buffer concentrations for the running buffer and sample buffer, respectively; |r and |s are the conductivities of the two buffers, which are in general proportional to cr and cs; and Es and Er are the electric field strengths in sample and running buffer zones. The electrophoretic velocity of each ionic species in the sample plug is proportional to the field strength, which leads to a rapid migration of anions to the back of the sample plug and cations to the front, assuming electro-osmotic flow towards the cathode. Once they reach the boundary to the high conductivity running buffer, the ions experience a sudden electric field drop, slow down and form a zone of concentrated sample ions at the end of the sample plug. Generally, the preconcentration efficiency increases with increasing k, but levels off for large k-values (\ 250, depending also on the separation parameters). Differences in conductivity and ionic strength between sample and running buffer lead to electro-osmotic flow being much higher in the sample than in the running buffer. This induces hydrodynamic flow effects as the system adjusts to ensure continuity of flow, and broadening of the sample plug as a result. These effects are described and analyzed in detail in Refs. [29,31,33].

Fig. 4. Mechanism of FAS. The electric field strength is concentrated in the low conductivity buffer plug and drives ions to the ends of the plug. Once they reach the high conductivity running buffer, they slow down and stack due to the lower field strength.

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Fig. 5. Signal gain as a function of relative conductivity for FAS on a microchip using 10 mM FITC-labeled Ser and Gly as sample (adapted from [31], reprinted with permission. Copyright 2001 Wiley – VCH).

FAS in 400 mm, volumetrically defined sample plugs on-chip yielded preconcentration efficiencies of up to 20-fold using fluorescently labeled amino acids as model analytes [31]. Injections were performed using a modified injection element, which allowed spatial confinement of the sample by two buffer streams from both sides without dilution of the sample plug. The advantages of volumetric definition are a high injection reproducibility (1.1% RSD peak height) and the absence of an electrophoretic sample bias. The presence and effect of hydrodynamic pressure on the stacking efficiency could be verified experimentally by CE (Fig. 5) and visually using fluorescence microscopy [31]. Higher preconcentration efficiencies require a larger amount of sample to be introduced into the system. Full-column stacking techniques using polarity reversal offer this possibility at the price of increased analysis time (1– 2 min typically). Li et

al. [32] used such a device to preconcentrate and focus trace level protein digests as preparation for quadrupole time-of-flight mass spectrometry (Fig. 6a). After the entire 70 nl channel between sample reservoir and on-chip nanoelectrospray emitter was filled with sample solution, the polarity was reversed to remove the sample matrix while the analyte stacked at the boundary to the running buffer. The progress of matrix removal could be followed by current monitoring, until only a small amount of buffer was left in the channel. Finally, the preconcentrated sample zone was pumped by EOF into the electrospray emitter for MS analysis. The concentration detection limit could be enhanced from three- to 50-fold, depending on the peptide. A different configuration was used by Lichtenberg et al., for full-column stacking of fluorescently labeled amino acids combined with on-chip CE for separation [31]. The design, depicted in

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Fig. 6b, consists of a 55– 170 mm long, folded stacking channel connected to a 40 mm long separation channel. In this way, high-efficiency stacking of a large sample volume could be combined with rapid separation in a short CE column. Filling of the stacking channel and the stacking itself were performed in 90 s, while the separation took only 35 s. The signal gain was found to be nearly linear with stacking channel lengths from 55 to 170 mm, yielding 60- to 95-fold increases.

Fig. 6. (a) Long-column stacking as preconcentration step before mass spectrometry: the channel between A and E is first filled completely with sample, then the sample is stacked by removing most of the sample buffer into reservoir C [32]. (b) Schematic chip layout of a column-coupled system with a long preconcentration channel (left) and a shorter separation channel (right). Both sections are connected via a 9 mm long common channel region [31] (reprinted with permission from [32,31]. Copyright 2000 and 2001 Wiley –VCH).

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3.2.2. Field-amplified injection Prior to the chip-based FAS examples described above, field-amplified injection (FAI) techniques on microchips were investigated by Jacobson and Ramsey [34] and Kutter et al. [16]. Fig. 7 presents the differences between FAS and FAI in a microchip format. While FAS works for both negatively and positively charged ions, FAI has an electrophoretic bias depending on the direction of EOF. In this case, positive ions are stacked at the sample–buffer interface during the injection process. FAI can be easily implemented using timebased, gated injection methods proposed by Jacobson et al. [35]. FAI yielded a concentration enhancement for dansylated amino acids of up to 13.8-fold at a k of 970. The efficiency was also investigated. Separations using stacked injection had efficiencies of 22000–29000 theoretical plates, which made them only 60 –70% as efficient as CE on the same devices using a volume-defined, pinched injection scheme. In combination with post-column complexation for detection, FAI was also used for the preconcentration of inorganic cations prior to CE [16], employing the same procedure as described above. The stacked injection mode improved the signal by a factor of 16, which corresponded to extrapolated detection limits of 18 ppb for calcium and 0.5 ppb for magnesium. The difficulty of controlling the precise location of the stacked sample zone in conventional chip designs has been overcome by a device recently presented in Ref. [36]. The electro-osmotic flow in these devices was eliminated by coating the channel walls using a 0.1% solution of poly(allylglycidyl ether-co-N,N-dimethylacrylamide) copolymer. Initially, all the channels were filled with a high-conductivity buffer by a pressure-driven pump system, except for the sample loading side-channel, which was filled with a lowconductivity buffer. Upon application of an electric field, the sample ions travel from the sample reservoir until they reach the stationary buffer concentration bound-ary and stack at this point. Once the desired amount of sample is collected, separation is carried out in the main channel of the chip. One of the major advantages of the stationary concentration boundary is that analyte collection can be carried out without dis-

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Fig. 7. While on-chip FAS allows preconcentration of both anions and cations in volumetrically defined plugs, field-amplified injection enriches only one ionic species at the front boundary of the injection plug.

placing the stacked plug, which led to a high preconcentration efficiency of more than 100-fold.

3.2.3. Stacking of neutral analytes Though relatively easy to implement, methods like FAS or FAI are applicable to charged analytes only. Methods for stacking neutral analytes for separation by micellar electrokinetic chromatography in fused silica capillaries have also been the subject of a number of recent papers. One approach, termed sweeping, involves the injection of an analyte zone into a column of separation buffer containing a pseudostationary phase (micelles). Upon application of the electric field, the micelles are swept through the analyte zone, collecting analyte as they go and resulting in a unique focusing effect. Charged species could also be preconcentrated in this way. When first described [37], preconcentration factors of several thousandfold were achieved. In a recent variation of sweeping, sensitivity increases approaching 1000000-fold have been reported [38]. Though not yet applied to microchips, it is clear that the use of this technique could be enormously advantageous, especially in conjunction with the integration of less sensitive absorbance or electrochemical detection methods into microfluidic devices.

Another method of neutral-species stacking in micellar CE involves addition of salt (e.g. NaCl) to the sample matrix to increase its conductivity to levels 2–3 times higher than the separation buffer [39]. The latter typically contains negatively charged micelles of sodium dodecyl sulphate (SDS) or sodium cholate. The result is a substantial drop in electric field strength at the sample plug–separation buffer interfaces, and a field amplification effect analogous to the one described above. In this case, it is the negatively charged micelles, with electrophoretic mobilities counter to the EOF that stack at the cathodic interface of the sample plug and the buffer. Analyte is collected and preconcentrated by the micelles as it is electroosmotically driven through this interfacial zone. The micelle zone is also effectively ‘locked’ in place up against the higher concentration chloride, until this latter zone has diffused sufficiently to let cholate anions enter. Hence, a sharp interface is maintained for analyte stacking. This type of stacking can be very effectively carried out during electrokinetic injection of sample, with plug lengths more than 100% that of the separation capillary [40]. This technique lends itself very well to chip-based separation, with 20-fold peak height improvement being observed for an 80 s

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injection of a solution containing 67 nM BODIPY [40].

3.2.4. Isotachophoresis for sample preconcentration and cleanup Isotachophoresis (ITP) extends the stacking concept of FAS to ternary buffer systems, and can be used both for sample preconcentration and as an analytical technique in its own right. The sample is injected between a leading and a terminating buffer electrolyte, LE and TE, respectively, with the condition that the mobility of the LE, vLE, is larger than the maximum v in the sample, and vTE is smaller than the minimum v in the sample. Upon application of the separation voltage, the sample constituents separate over a given time into distinct zones located between LE and TE in order of descending mobilities. Once this steady state is reached, the boundaries between sample zones are very sharp, due to a self-focusing mechanism governed by the Kohlrausch regulating function [41,42]. The concentration in the sample zones is equal to the concentration of the leading buffer. Therefore, a preconcentration or dilution can be tailored by adapting the LE composition. Kaniansky et al. realized various ITP configurations on a poly(methyl methacrylate) (PMMA) chip with two coupled separation columns and on-chip conductivity detectors [43]. One application involving a combination of sample pretreatment by ITP and separation by CE involved the analysis of low concentration constituents (10 mM each of nitrite, phosphate and fluoride) in a model mixture containing high background concentrations of ions like chloride (600 mM) and sulfate (800 mM). A sample fraction containing the micro-constituents could be extracted from the steady-state ITP zone system and electrophoretically transferred into the second column for CE separation. The fabrication of these PMMA chips with integrated electrodes for conductivity detection is explained in more detail in Ref. [44], where performance is demonstrated by separation of a sample containing organic acids. Another polymer microfluidic device for ITP separation of metal cations, this one formed in silicone rubber by casting, has recently been re-

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ported [45]. Here too, detection was performed using an integrated single-electrode conductivity detector, indicating the utility of this not-so-sensitive detection technique when used together with a preconcentration method like ITP. The separation of a sample containing a mixture of the four metal ions lithium, lanthanum, dysprosium and ytterbium was reproducibly achieved using these devices. The miniaturized separations were achieved in under 600 s, which is less than half the time taken for capillary-scale separations.

3.2.5. Solid-phase extraction techniques In general, solid-phase extraction (SPE) refers to processes where the analyte is retained by an appropriate stationary phase and subsequently eluted in a more concentrated form. For this discussion, SPE is considered in a broader sense, with examples involving isolation of analyte from an interfering sample matrix, as well as preconcentration, being described. A perhaps somewhat arbitrary division of SPE methods into two categories, termed non-selective and selective, has been made. This reflects the degree of selectivity of the surface layers used for extraction in the different cases. In the former case, surfaces adsorbing a wide range of compounds were used, whereas in the latter, selectivity was imparted to the surface through immobilization of very biospecific species. 3.2.5.1. Non-selecti6e SPE. Two main strategies for this type of on-chip SPE appear in the literature, using either immobilized stationary phases on channel walls or channels packed with coated beads. The first method has been more prominent to date, as it is compatible with standard microfluidic systems. Kutter et al. [46] demonstrated SPE of a neutral coumarin dye, C460, in a simple glass microchip. The fluidic network consisted of a cross configuration, but with a side channel containing an additional branch and reservoir, to be able to switch between enrichment and elution buffer. To coat the channel walls with a C18 stationary phase, the chip was filled with octadecyltrimethoxysilane in dried toluene at room temperature for 24 h [47]. For enrichment, 8.7 nM sample was electrokinetically pumped through the

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channel at a field strength of 360 V cm − 1 in a pH 8.5 sodium tetraborate buffer containing 15% (v/ v) acetonitrile. After an enrichment time of up to 5 min, the extracted dye was eluted using the same buffer, but containing 60% acetonitrile. The gain in concentration achieved was up to 80-fold, and increased rapidly with enrichment time up to ca. 160 s, after which the enhancement leveled off. For a 160 s enrichment time, the enriched volume was 25.6 nl, while the elution volume was only 0.48 nl, assuming a 3 s wide elution peak. This led to a predicted 50-fold concentration increase for rectangular plugs, which, when corrected for a Gaussian profile, meant an estimated 80-fold increase, in agreement with the experimental data. The capacity of an SPE column strongly depends on the surface area available for binding. In order to increase this parameter with respect to open-channel SPE, Oleschuk et al. [48,49] created a packed bed of octadecylsilane (ODS)-coated silica beads (1.5–4 mm) in a microfluidic network. To do so, a 200 mm long, 580 mm wide and 10 mm deep cavity was etched into the glass, with its in and outlet channels obstructed by 9 mm high weirs to leave a 1 mm gap for solution introduction (Fig. 8). Beads greater than 1 mm in diameter could thus be flushed into the cavity through a third, 30 mm wide side channel, and be retained behind the weirs. By means of the side channel, the beads could also be removed from the bed for replacement by new packing material. A 1 nM solution of BODIPY dye was used as test substance and flushed through the bed in an aqueous buffer for times varying from 2 to 10 min. After washing the excess sample out with buffer, the captured BODIPY was eluted in acetonitrile. Preconcentration factors of 80 up to 500 could be achieved in this configuration. Owing to the increased effort needed for fabrication of packed bead beds on-chip, conventional, miniaturized, off-chip SPE has been coupled to microchips. A C18-SPE cartridge mounted between a microchip and an electrospray needle was used for MS protein identification, where the microchip served as nanoflow solvent gradient source [50]. Li et al. have compared the performance of a C18-SPE membrane preconcentrator for protein digests with field-amplification stack-

ing (FAS) as described before [32]. Preliminary results indicated limits of detection in the nanomolar range, which positions SPE in the same efficiency range as full-column FAS.

3.2.5.2. Non-selecti6e SPE of DNA. After extraction from the living cell, a purification step is often necessary to isolate the nucleic acids from other interferents, such as proteins or metal complexes. If the purification process is based on binding of the DNA and RNA to a stationary phase, it can also include a preconcentration effect. Many commercial nucleic acid purification kits are based on the fact that DNA can be effectively extracted by binding to silica particles. An established, small-scale purification method known as the Boom method utilizes a high concentration of guanidinium thiocyanate (GuSCN) as chaotropic agent, which not only improves DNA binding but also lyses cellular material and inactivates nucleases [51]. This procedure was

Fig. 8. (a) This bead-packing device takes advantage of two weirs at the in- and outlet of the SPE cavity, which retain the ODS-coated beads. The gap between weir and cavity cover is 1 mm, while the beads have a diameter of 1.5 – 4 mm. (b) The SPE cavity is filled via the bead introduction channel 3, while the weirs stop beads from entering the in- or outlet channel 1 and 2 (reprinted with permission from [49]. Copyright 2000 Kluwer Academic Publishers).

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Fig. 9. (a) Device layout for the CE chip with DNA preconcentration at a porous membrane at the injection junction. For preconcentration, an electric field of 1 kV is applied between sample and side channel reservoirs for 1 – 2 min to cause DNA to migrate and collect on the junction [54]. (b) Electropherograms of a DNA PCR marker after different preconcentration times on a similar device [53] (reprinted with permission from [54,53]. Copyright 2000 and 1999 American Chemical Society).

transferred by Christel et al., to a deep-reactiveion-etched silicon microstructure, which used a regular array of 200 mm high, 18 mm diameter posts rather than silica beads to create a higher surface area for DNA extraction [52]. After oxidation of the silicon and anodic bonding to a glass plate, the purification device could be used in a flow-through fashion. The purification protocol has three steps: (1) DNA collection in a chaotropic salt solution; (2) washing using an ethanol-based solution; and (3) elution in water or suitable buffer. For low-concentration DNA samples (less than 105 copies of l DNA), a ten-fold concentration effect and a 50% capture efficiency were observed. Recently, a new way of DNA sample preconcentration was introduced in the form of integrated porous membrane structures [53,54]. These

planar membranes allowed DNA fragments to be concentrated up to 100-fold before separation. The device structure, depicted in Fig. 9a, consisting of a typical microchip CE layout with an additional U-shaped side channel, which is close to the injection element but not in direct contact (a 3–12 mm wide gap was maintained between the channels). Bonding of the structured glass wafer to the cover plate is achieved at low temperatures by spin-coating a thin layer of diluted potassium or sodium silicate solution onto the coverplate as adhesive. Due to its porosity, this layer also acts as a semipermeable membrane between the Ushaped side channel and the main channel in the region of the injection junction. Ionic current can flow over the junction, while larger molecules are retained. Preconcentration can be achieved by applying a voltage of 1 kV between the sample

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and side channel reservoirs for a given time, which causes the analyte to migrate to the membrane region. Fig. 9b shows three successive electropherograms obtained for different preconcentration times of a DNAPCR marker, indicating a non-linear relationship between signal intensity and time. In a follow-up study, the porous membrane preconcentrator was used after on-chip PCR to reduce the number of thermal cycles necessary for product detection down to ten, leading to a total analysis time of less than 20 min [54]. Along the same lines, Lin et al. [55] recently developed a PMMA-based microchip with an inchannel metal electrode for DNA preconcentration without the use of a porous membrane.

3.2.5.3. Selecti6e SPE. Selective extraction and preconcentration of biological molecules onto a surface bearing immobilized antibodies, DNA oligomers, or other molecular recognition elements can also be considered a form of SPE. For example, antibodies may be purified by reaction with protein-coated beads. In this case, antibodies showing a high affinity for the probe molecules are retained on the particles, while remaining species are washed away. They can then be subsequently released for further use as reagents in a bioanalysis. The preconcentration aspect of a biochemical SPE process is exploited in heterogeneous assays. Though these methods are not considered sample pretreatment methods per se, they do owe their excellent sensitivity (nM to pM) to the high affinity of bound probe species, usually antibodies, for the analyte in solution. Given a suitable amount of time, this can lead to significant preconcentration of analyte on the surface, particularly from dilute samples. This section will discuss some examples of selective SPE carried out in microfluidic devices. Several examples of heterogeneous immunoassay on chip will also be considered in this context, since the platforms developed for these analyses could, with some modification, be also used for sample pretreatment prior to analysis. Reports of microfluidic devices for selective extraction and isolation of biological molecules are still relatively uncommon. One recent, notable example is that of Jiang and Harrison, who re-

ported the isolation of mRNA from samples using commercially available, paramagnetic oligo-dT beads [56]. These polystyrene beads, which are less than 3 mm in diameter, are prepared with covalently attached chains of deoxythymidylate (poly(T)). This makes them ideal for rapid isolation of RNA from cell extracts, since most RNAs have a poly(A) tail. Beads and sample introduced from two arms of a Y-junction could be rapidly mixed by diffusion. Reacted beads could then be magnetically trapped while sample was flushed away, then released again. It was estimated that as much as 34 ng of mRNA could be retained from 10 mg of total RNA in a sample. Fan et al. attached different DNA sample fragments to the poly(T) chain of oligo-dT25 (25 T’s) beads, and tested their reactivity with a fluorescent DNA probe [57]. Modified particles were held in place with a magnet in a 750 mm long microchannel segment to form a 2 nl plug, and perfused with the probe-containing solution using pressuredriven flow. After hybridization, denaturation by on-chip heaters was used to remove the bound molecules again. Hybridization was observed to take place enormously rapidly, in just a few seconds, in stark contrast to the 3–18 h required when dense arrays of immobilized oligomers are used as probes. It was concluded that the facilitated interaction of probe and sample, due to rapid delivery of probe to the beads, was in part responsible for this increased hybridization rate. The small sample volume involved also meant that a higher concentration of probe could be used without concern for cost or supply, so that hybridization could be driven much more rapidly to completion. This paper described an analytical approach developed with gene expression analysis in mind. However, it is mentioned in the sample pretreatment context as well, because it so clearly demonstrates the unique advantages microfluidics can bring to solid-phase extraction and isolation processes with respect to speed and ease of smallvolume liquid handling. In the above examples, conventional magnets were used to manipulate beads. Recently, Choi et al. successfully integrated a planar electromagnet into a flow chamber to retain magnetic beads for selective sampling of biomolecules from cell extracts [58].

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Several examples of integrated heterogeneous assays on chip point the way to selective sampling devices for other applications, and are therefore mentioned here. Fig. 10 shows the fluorescent signal acquired for one complete, protein A-rabbit immunoglobulin G (Cy5-labeled) binding assay in an open channel [59]. Protein A was immobilized on the silanized glass walls of a 165 pl reactor channel segment. Cy5-rIgG was first incubated with the protein A by flowing this solution through the reactor for a period of 200 s, then washed away. Bound Cy5-rIgG was then eluted with a glycine–HCl (pH 2) buffer, to yield a sharp peak in the fluorescence trace. Comparison of this peak with the signal obtained during incubation indicated that a signal gain of up to 30-fold for an incubation time of 200 s could be observed. Sato et al. have performed the analysis of secretory human immunoglobulin A using a packed bed of 45 mm diameter polystyrene beads in a microchip [60]. The target was adsorbed on the bead surface, which was then reacted with colloidal-gold-conjugated anti-s-IgA antibody and detected by a thermal lens microscope. Analysis times for this analyte could be reduced from 24 h to less than 1 h. The same system was also used to analyze a carcinoembryonic antigen [61].

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3.3. Bioanalytical sample pretreatment 3.3.1. Polymerase chain reaction Since its introduction by Mullis et al. [62], the PCR has become a key technology for genomics. This reaction synthesizes DNA from a starting template, a DNA oligomer, using a cycle of polymerase enzyme reactions. The concentration of DNA increases exponentially after each cycle, so that typically after 20 to 30 cycles, there is enough DNA available for further analysis by conventional gel electrophoresis. The key to this reaction is repeated and controlled temperature cycling between a denaturation temperature ( 90– 95 °C), an extension temperature ( 70 °C) at which the polymerization occurs, and an annealing temperature (50 °C) at which strands rehybridize. Conventionally, PCR is done in benchtop instruments, whose large thermal mass limits the speed at which thermal cycling can be carried out. Microchip PCR therefore offers an ideal approach for decreasing the time PCR takes by providing devices with low thermal inertia, reducing typical cycle times from several minutes to 30 s or less. The proof-of-principle for PCR in silicon microstructures was initially presented by Northrup et al. in 1993 [63] and Wilding et al. in 1994 [64].

Fig. 10. Photomultiplier (PMT) response for a complete on-chip immunoassay cycle yielding a sample preconcentration of ca. 20-fold in this case (Cy5-rIgG concentration is 1.5 mM) (reprinted with permission from [59]. Copyright 2001 American Chemical Society).

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In both cases, the chip contained an etched reaction chamber with a volume on the order of several microliters and an in- and outlet. The devices were sealed using a Pyrex glass plate and anodic bonding. In the former case, a polysilicon heater was integrated into the device, allowing thermal cycling at rates 2– 10 times faster than conventional instruments, with comparable yields. In the work described in Ref. [64], heating and cooling was achieved by an external thermoelectric device on a copper block, yielding comparatively short cycling times in the order of 3 min. The device was successfully used to amplify a 500 bp segment of bacteriophage l DNA. A concern when using silicon as substrate for high surface-to-volume ratio reactors is the material compatibility with the reagents used. Different silicon-based surface passivation layers for PCR chambers were evaluated in Ref. [65]. It was found that native Si and Si3N4 caused a high failure rate, while silanization followed by a polymer coating showed good but inconsistent amplification, and SiO2 performed best. Similarly, alternatives to silanization for passivation of surfaces in glass PCR micro-chambers have been explored by Giordano et al. [66]. These researchers studied the surface interaction properties of the additives polyethylene glycol, polyvinylpyrrolidone, and hydroxyethylcellulose, all of which act as dynamic coatings, and epoxy (poly)dimethylacrylamide, a compound that adsorbs to glass. Analysis of PCR products generated in the microchips under different conditions revealed that these coating agents adequately passivated chamber surfaces. Optimization of PCR reagent mixes for silicon-based devices has been published by Taylor et al. [67]. Since the initial silicon-based PCR chambers appeared, a number of technological developments have been reported for various applications. A stand-alone version of the silicon PCR reactor in Ref. [63] has been refined further [68] and a cartridge device using disposable polypropylene reaction tubes is commercially available [69]. More recently, another PCR chip in silicon, with integrated Pt temperature sensors and heaters, was reported [70]. This device could control temperature very accurately in real-time

to within 9 0.025 °C. Again, a surface treatment involving either silanization followed by coating by polyadenylic acid, or oxidation of the chamber walls, was required to obtain efficient PCR amplification. The device was tested using four traditional Chinese medicine genes as templates, and found to yield DNA concentrations of 27 ng ml − 1 or more, sufficient for further analysis. Probably the most technologically advanced realization of an integrated DNA analysis device has been developed at the University of Michigan [71,72]. The 47-by-5 mm large silicon chip contains a sample loading stage with a drop-volume-metering feature, a mixer, a thermal reactor for amplification, a gel loading stage, and a gel electrophoresis channel with integrated optical detection system. Fluid handling is achieved by a set of hydrophobic vents and external air pressure, which drives single droplets precisely through the channel system. Most examples of integrated PCR depend on a direct contact between the PCR chamber and the heating source to achieve fast thermal cycling. To circumvent the technical issues associated with contact methods, Oda et al. reported the use of a non-contact thermal cycling method, using an inexpensive infrared radiation source for heating and a solenoid-gated compressed air source for cooling [73]. Cycle times as fast as 17 s could be achieved in glass micro-chambers. In a more recent polymer embodiment of the chip and using infrared-mediated temperature control, this group has achieved adequate amounts of PCR product after only 15 cycles, for a total amplification time of only 240 s [74]. In 1996, the first integration of a planar CE device with a microchip-based PCR reactor was reported [75]. The device consisted of the hybrid assembly of the planar CE chip for DNA separation together with a polypropylene reaction chamber in a polysilicon heating mantle. The PCR reactor served simultaneously for DNA amplification and subsequently as the sample reservoir for CE analysis. The low thermal inertia of the integrated heating system enabled heating rates of 10 °C s − 1, while passive cooling took place at 2.5 °C s − 1, for an overall cycle time of 30 s. The

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Fig. 11. An array of eight coupled PCR-CE systems. The sample is loaded via a fluidic bus A, through the open valve B into the PCR chamber C, where it stops because of the hydrophobic vent D. After the amplification is finished, the sample is loaded onto the separation channel and sized [76] (reprinted from Ref. [76] with permission. Copyright 2001 Elsevier Science).

approach was further refined through integration of 280 nl PCR chambers directly into the glass chip, in the same plane as the CE channels, as depicted in Fig. 11 [76,77]. The device contains microfluidic valves and hydrophobic vents to immobilize the sample in the reaction chamber during PCR, which is driven using a resistive heater clamped onto the chip. The device contains eight parallel amplification/separation channels on a single wafer. In this system, single-molecule DNA amplification and analysis has been achieved [77].

Another example for on-chip PCR combined with CE is a microchip device for cell lysis, multiplex PCR amplification and electrophoretic sizing in conventional glass technology [78]. In this case, PCR was effected in one of the sample reservoirs, with the whole chip being cycled in a conventional thermocycler. A similar system was used to run multiple-sample PCR amplification on a chip, followed by sizing of the PCR products using onchip CE [79]. Recently, an arrangement of two Peltier thermoelectric elements was used to re-

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place the benchtop thermocycler, yielding cycle times of less than 1 min. A DNA preconcentrator was integrated on-chip by a micromachined frit (see Section 3.2.5.2) to reduce the number of cycles necessary for analysis (total reaction time was less than 20 min) [54]. A combined PCR – CE hybrid device, formed from poly(dimethylsiloxane) (PDMS) and glass layers, was recently reported for a gene amplification application [80]. PCR was successfully demonstrated in this chip using a 500 bp l DNA target, and the products subsequently analyzed. The use of PDMS, an inexpensive material with excellent optical characteristics, could make this a route to single-use devices. A distinctly different approach to on-chip PCR was introduced by Kopp et al. [81]. In this case, the chip is in contact with three side-by-side copper blocks, which provide three distinct constanttemperature zones. Instead of repetitively heating and cooling a sample contained in a microreactor, the sample plug is pumped at a constant velocity through a meander-like glass channel from one temperature zone to the other (Fig. 12). In this way, the effect of thermal inertia is eliminated.

Moreover, a large number of sample plugs separated by a spacer liquid can be amplified in parallel. Heating and cooling times were less than 100 ms, and total amplification times between 1.5 and 18.7 min were obtained, simply by changing the flow rate. The amplification gain decreased with decreased cycle time, but even for short, 12 s cycles, a good fluorescence signal could be obtained after 20 cycles.

3.3.2. Enzymatic digestion: DNA Restriction enzymes are one of the most powerful tools for recombinant DNA analysis. The use of microfabricated devices to combine this sophisticated biochemical sample pretreatment process with CE separation has been demonstrated by Jacobson and Ramsey for DNA restriction fragment analysis [82]. On-chip, it took only 5 min at room temperature to digest the plasmid pBR322 using the enzyme HinfI and to size it using a polymeric sieving matrix. Fig. 13 shows the digestion procedure that takes place within a 0.7 nl reaction chamber on a glass-based chip device. Electro-osmotic flow was minimized by covalent immobilization of linear polyacrylamide on the

Fig. 12. A continuous-flow PCR chip as proposed by Kopp et al. [81]. Copper block A is held at 95 °C for denaturation, B at 77 °C for extension and C at 60 °C for annealing. The sample is transported by pressure driven flow through the meander glass channel (reprinted from Ref. [76] with permission. Copyright 1998 American Association for the Advancement of Science).

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Fig. 13. Operation procedure for DNA digestion and fragment sizing: (a) the reaction chamber is filled with enzyme and DNA, the excess being removed into the Waste 1 reservoir; (b) after the digestion period, the DNA fragments are moved by a time-controlled, gated injection into the separation column; voltages are given relative to the voltage HV at the Waste 2 reservoir, field strengths were varied between 190 and 690 V cm − 1 (diagram adapted from [82], reprinted with permission. Copyright 1996 American Chemical Society).

channel walls, and hydroxyethyl cellulose was used as sieving medium. Initially, the reaction chamber was filled with electromigration towards the Waste 1 reservoir of a mixture of the DNA sample (125 ng ml − 1) and the enzyme solution (4 units ml − 1). This reservoir was filled with a buffer mix containing cations necessary for the digestion, such as Mg2 + , which also migrate towards the reaction chamber. Digestion could be performed either in a transient, continuous flow mode, yielding a reaction time of 9 s using the voltages indicated in Fig. 13, or in a static mode with all voltages switched off for a given digestion time. Peak height increased tenfold by increasing the digestion period from 9 to 129 s, but no further increase could be observed for longer reaction times. This effect could not be completely explained, though it was postulated that it might be due to completion of the digestion or loss of enzyme to the walls of the reaction chamber because of its high surface-to-volume ratio. However, experiments showed that adsorption losses of DNA were not so significant.

3.3.3. Enzymatic digestion: proteins Several devices reported recently embody a high degree of functionality, integrating protein digestion with subsequent analysis. One such device for combined digestion, separation and post-column labeling of proteins and peptides using naphtha-

lene-2,3-dicarboxaldehyde is described in Ref. [83]. A tryptic digest of oxidized insulin B-chain took 15 min to go almost to completion in a heated channel segment, followed by labeling and electrophoretic separation. The integration of proteolytic digestion on a microchip with mass spectrometric detection was demonstrated by Xue et al. [84] for the analysis of peptides generated by tryptic digestion of melittin. The digestion was performed in the sample reservoir to be able to monitor the process over time by repetitive MS analysis. Harrison et al. [85] recently introduced a chip design featuring a 15 mm long, 800 mm wide, and 150 mm deep bed of packed, 40–60 mm, trypsincoated beads for on-chip protein digestion prior to introduction to an electrospray mass spectrometry interface. The device also included a 45 mm long separation channel. The rapid digestion, separation, and identification of proteins were demonstrated for melittin, cytochrome c, and bovine serum albumin. Generally, digestions took from 3 to 6 min. However, melittin was consumed within 5 s, in sharp contrast to the 10–15 min required in a conventional cuvette. The use of an integrated digestion bed together with hydrodynamic flow effectively increased the ratio of trypsin to sample, to overcome the kinetic limitations often associated with digestion of small amounts (picomoles) of protein. Again, this is an

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example of the use of microfluidics to generate optimal conditions for a chemical reaction in a unique way not possible in macroscopic systems.

3.3.4. Other techniques In order to improve both speed and reliability of DNA binding assays, Nanogen has developed a 1 cm2 chip platform that allows the sample to be moved and removed from specific binding sites in a controlled fashion [86,87]. To do so, a square array of 25– 10000 metal electrode pads, with sizes ranging from 80 down to 30 mm, is fabricated on a silicon chip using standard microelectronics processing technology, together with arrays of auxiliary and counter electrodes. The array is covered with an agarose-based permeation layer that has different capture probes anchored to it by streptavidin– biotin attachment. To attract the sample DNA to a specific binding site, the electrode at that location is positively biased with respect to the adjoining counter electrode. Negatively charged DNA molecules are thus attracted to the site and exhibit increased binding rates because of their preconcentration at the electrode surface. Additionally, by inverting the polarity, an ‘electrical wash’ of the electrode surface is initiated, which removes with increasing electric field strengths unbound target DNA, non-specific DNA and partially hybridized target DNA, in that order. Recently, DNA amplification at a binding site could also be demonstrated [88]. 3.4. Separating sample from sample matrix 3.4.1. Physical particle filtering and retention by integrated flow restrictions Clogging of channels due to particle contamination is a common problem when working with miniaturized analysis systems. The small crosssections of microfluidic channel networks require filtering of most liquid samples prior to analysis. The laboratory answer is pre-filtering of the solutions using syringe filters. However, this method is incompatible with the ultimate goal of mTAS, which is autonomous operation and integration of the complete analysis process into a single device. An overview of the size ranges

relevant to filtering and dialysis systems is given in Fig. 14. Interestingly, the same types of microstructures used to prevent particles from entering into a microfluidic system can also be deployed for the opposite purpose, namely to retain particles like chemically coated polymer beads or biological cells in a reaction volume. The examples given in this section are taken from both filtering and retention applications, and give an overview of some of the advanced fabrication technologies available for silicon-based mTAS. A straightforward approach to the filtering problem is the implementation of flow restrictions in the flow path of the channel network. Thus, particles having a cross-section larger than the gap between the barriers are retained behind, while solution can freely pass through the restriction. These so-called axial percolation filters can be fabricated by simple one-mask photolithography processes using anisotropic reactive ion etching or conventional wet chemical etching [89]. In this example, white blood cells could be efficiently isolated by filtering them through a network of silicon pillars with a gap of 5 mm. Due to the elasticity of the cells, the pillars had to be rather long in the flow direction (175 mm) to retain the cells, since the cells deformed and easily slipped through the channels between shorter pillars. Due to the under-etching of the mask, wet chemical etching turned out to be less suitable for the fabrication of such devices, especially as structures with rather small openings are required. Recently, deep reactiveion etching (DRIE) of silicon has been used to work around the problem of low aspect ratio features [90]. Arrays of 50 mm high posts (3 mm× 10 mm) with spacings as close as 2 mm were used to capture polystyrene beads with a diameter of 5.5 mm (Fig. 15). The aim here was not to remove particles from the analyte solution, but rather to pack the complete filter volume with a stationary phase to perform a chemical reaction. Recently, a novel bead retention concept, not involving physical barriers, has been introduced, which allows packing of bead beds with less stringent requirements for fabrication technology [91].

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The design strategies mentioned before are currently limited to filter cut-off sizes of about 1 mm, which corresponds to the resolution limit of conventional photolithography. However, to achieve results comparable to standard syringe filters, the feature size of the elements composing the flow restriction network has to be less than 0.5 mm. Microelectronic foundries have long been working in the sub-micrometer feature size range for planar devices (etching depthB0.5 mm). However, the fabrication of microfluidic structures of that size with micrometer depths is extremely difficult, because of the poor depth-to-width ratios of most etching processes. To overcome this problem, several groups have proposed sacrificial layer (SL) techniques to create tortuous flow paths with restrictions in the range of tens of nanometers for efficient filtering [92– 95]. Here, the smallest feature size is not directly

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defined by lithography, but rather by deposition of a sandwiched, thin, intermediate material layer. This is subsequently selectively removed, and the gap that remains acts as a filter ‘pore’. Deposition techniques like chemical or physical vapor deposition (CVD and PVD) can be controlled very precisely in the nanometer range, so that filters with pore sizes of down to 10 nm were fabricated [93]. Fig. 16 shows the schematic fabrication procedure for an SL filter as proposed by Kittilsland and Stemme [92]. The filter principle is based on two planes perforated by a regular hole pattern, which are offset by half of the hole diameter. A thin gap between the two planes allows liquid flow from one side of this membrane structure to the other. To fabricate this structure, a thin layer of thermal silicon oxide, covered by polysilicon, serves as SL. Access holes are etched into the

Fig. 14. Size ranges for filtering and dialysis applications (adapted from ‘The Filtration Spectrum’, Osmonics Inc., Minnetonka, MN, reprinted with permission. Copyright Osmonics, Inc.).

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needs to be assessed, as there are various alternatives requiring much less technological effort. A different approach to the filtering problem was adopted by He et al., who decided to work with a lateral percolation filter [98]. In contrast to the axial type described above, filtering action is achieved here by retention of the particles in a bed of posts, through which the fluid flows laterally. In the given example, an array of posts 1.5 mm wide and 10 mm high was machined into a quartz wafer by reactive ion etching at the bottom of the reservoirs leading into the microchannels. Liquid Fig. 15. A flow-through reactor featuring a filter structure composed of a row of pillars fabricated by deep reactive-ion etching. The pillar dimensions are 3 × 10 × 50 mm3 with a spacing of 2 mm [90] (reprinted from [90] with permission. Copyright 2000 Elsevier Science).

polysilicon layer by dry etching, followed by removal of the sacrificial layer to open the filtration pores. Subsequently, holes are etched into the backside of the filter by dopant-selective etching in potassium hydroxide solution [96]. The final structure has a 1.5 mm thick filter membrane with inlet and outlet holes in the range of 10 mm. The cut-off size of the filter can be tailored by changing the SL thickness, which is 50 nm in the example given above. A different approach was taken by Kuiper et al. [97] to machine an array of small holes with diameters down to 65 nm and a pitch of 200 nm into thin silicon nitride membranes about 100 nm thick. The authors used laser-interference photolithography to create a pattern of parallel fringes on the wafer. A thin layer of photosensitive resist deposited on the wafer served as mask for the subsequent etching of filter pores into the membrane. By adjusting the exposure dose to half of the necessary dose and superimposing two fringe patterns at a 90° angle, all areas of the resist where two fringes crossed received a double dose of light and were thereby polymerized. After development, a regular array of square posts remained. Despite the many interesting features of the filter structures above, the practicality of their fabrication using photolithographic processes still

Fig. 16. (a) Schematic principle of a sub-micron sacrificial layer filter. (b) A thin layer of silicon dioxide acts as spacer for the flow restriction opening. It is removed (or sacrificed) at the end of the fabrication procedure. (c) Close-up scanning electron micrograph of the final structure, showing the perforated polysilicon membrane on top and the hemispheric bulk silicon membrane below [92] (reprinted from [92] with permission. Copyright 1990 Elsevier Science).

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Fig. 17. (a) Schematic layout of the planar filter structure of Brody et al. (adapted from [102]). A particle-free fluid is pushed by the source stream through the barrier while particles are washed out through the larger outlet channel. (b) The cross-section shows the two parallel channels fabricated by anisotropic etching of silicon, with a thinner barrier gap covered by a glass lid (reprinted and adapted from [102] with permission. Copyright 1996 Elsevier Science).

entering the system from above would pass the network of channels between the posts and then exit laterally into the channel. Particles larger than the 1.5 mm distance between the posts would be held back without clogging the filter, as many other surrounding channels around still allow free liquid flow. However, both the distance between the posts as well as the filter depth are difficult to reduce, due to limitations of the etching process used.

3.4.2. Filtering by diffusion in laminar flows A unique feature of microfluidic systems is the dominance of laminar flow conditions at the flow rates normally used (nl s − 1 – ml s − 1), due to the small dimensions of the channels. An interesting quality of laminar flow is that material transport between two adjacent streamlines only takes place by diffusion, and not by turbulent mixing as it is done in a test tube. A characteristic value to describe the fluidic behavior in a channel is the Reynolds number, Re, a non-dimensional parameter relating inertial and viscous forces of the liquid. z Re =U( Dh p

(2)

where U( is the average fluid velocity, Dh is the hydraulic diameter, z is the fluid density, and p is the fluid dynamic viscosity.

This parameter can be used to predict whether a flow in a given duct is laminar or turbulent. For non-circular flow paths, laminar flow prevails at Re B2000, whereas turbulent flow exists at Re \ 3000 [99]. In fact, in many microfluidic cases Re is even well below 1 [100]. For microchannels of various cross-sectional geometries, Dh is 4A/P, where A is the cross-sectional area and P is the wetted perimeter of the channel. For water at room temperature, the kinematic viscosity p/z is 1.007×10 − 6 m2 s − 1, which leads to an Re of 0.02 for a 20 mm diameter circular channel at a flow velocity of 1 mm s − 1. Diffusional filtering can be achieved if laminarly flowing sample and buffer streams are brought in contact for a given time. This contact time can be tuned such that lighter molecules having a high mobility can cross over into the buffer stream, while heavier particles remain in the source flow and are finally washed out of the system. To exemplify this, it should be noted that for instance a small organic dye molecule with a molecular weight of 330 Da takes roughly 0.2 s to diffuse 10 mm, whereas a polymer bead of 0.5 mm diameter travels the same distance in 200 s [101]. Devices based on this type of filtering, but still having a physical barrier, were first introduced in 1996 by Brody et al. [102]. In the first device design, a pressure-driven, laminar, source fluid

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stream flowed along a thin barrier over a certain distance into an outlet reservoir (Fig. 17). Particles larger than the barrier gap did not pass it and only filtered fluid was pushed through the constriction into the outlet channel for the filtrate. In practice, the filter structure was fabricated using a three-mask process based on anisotropic etching in EDP (ethylene diamine, pyrocatecol and water) solution. Fluidic connections were first etched through the wafer from the backside using silicon dioxide as mask, then the fluidic channels were formed on the front side, and finally the 1 mm deep barrier gap was etched. To cover the channels, a piece of Pyrex glass was anodically bonded to the silicon chip. The device was successfully tested with 16 and 2.6 mm diameter fluorescent spheres, the first being filtered completely while a few of the latter could cross the barrier. Brody and Yager from the same group extended the idea to a filter system that does not need a physical separation barrier and fully relies on laminar flow [103]. As Fig. 18 shows, an H-shaped microchannel system is fabricated to be able to bring two laminar flows in contact over a certain distance and separate both streams shortly after. One stream contains the source liquid while the other serves as receiving stream for the particle-free sample. As long as laminar flow is assured, mass transport from one stream to the other is not dependent on fluid motion but only on diffusion. Later, diffusion-based systems were also used for direct, basically calibration-free albumin assays [104,105].

3.4.3. Microdialysis Dialysis is a long-established technique for cleanup of samples of both biological and non-biological origin. It involves the use of a semipermeable membrane to separate the sample stream from a perfusing liquid, also known as perfusate or microdialysate, a solution of well-defined composition. Exchange of species between these liquids is strictly controlled by the presence of the membrane, whose pore structure will allow transport of molecules only up to a certain size or molecular weight. Molecules larger than the socalled molecular weight cut-off (MWCO) will not be able to cross over into the perfusing liquid. However, species having a MW less than the MWCO can diffuse across the membrane, a process which is driven by the concentration gradient between the two streams. Dialysis membranes having MWCOs ranging from 100 Da to 300 kDa are commercially available, and are often made from materials such as cellulose ester or regenerated cellulose. Microdialysis can work in two ways as a sample cleanup method. In the first, the analyte of interest can be extracted from a complex matrix into the clean perfusate, which is then subsequently analyzed downstream. This has been effectively applied in real-time sampling of physiological fluids in animals and humans, where analysis is facilitated by first transferring the analyte into perfusate and then transporting to an ex vivo analytical device or system [106,107]. Biopro-

Fig. 18. The H-filter is based on two laminar streams, which are in contact for a short time to allow highly mobile analyte to cross over from one stream to the other while heavier particles remain in the source stream [103] (reprinted from [103] with permission. Copyright 1997 Elsevier Science).

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cesses and environmental systems as dirty as waste water may also be sampled in this way [108]. Alternatively, small interfering contaminants can be removed from a sample stream, which can then be more effectively analyzed. An example of dialysis for this type of application is the removal of high levels of salt from samples before analysis by mass spectrometry. The coupling of a microdialysis probe with a micromachined flow manifold, containing an integrated biosensor array, for real-time monitoring in a clinical setting of glucose and lactate in blood has been reported [109,110]. The use of biosensors in this instance dictated a sample pretreatment step producing a clean, protein-free sample. This is because sensor lifetimes are generally severely compromised when exposed to heterogeneous, physiological fluids like blood, due to fouling of the surface sensing layer. High analyte recoveries in microdialysis sampling require very low flow rates, on the order of 1 ml min − 1 or even nl min − 1. As this was the case here as well, it was decided to work with a microflow system, to avoid as much as possible the dead volumes which lead to slowed sample transport and longer analysis times. The microdialysis probe used was a small-scale, extracorporeal shunt circuit, described in more detail in Refs. [109,110], which provided 100% sampling efficiency of glucose and lactate from blood. During in vivo testing of dogs, the system demonstrated a response time of less than 5 min, thus effectively allowing real-time monitoring of the analytes of interest [110]. The use of microtechnology to produce low dead-volume connections between a needle-like dialysis probe and a microchip incorporating a chloride sensor for direct measurements in subcutaneous tissue or blood has also been proposed by Bo¨ hm et al. [111]. The dialysis probe was fabricated by slipping a smaller fused silica-capillary (125 mm OD) into a larger one (320 mm ID) and sealing the tip using a small tube made from a suitable dialysis membrane (300 mm OD) (Fig. 19). The perfusion liquid passed through the outer tube into the tip region, taking up the sample constituents of interest and flowing via the inner capillary into the analysis device. Chloride speciation was done using a Ag/AgCl electrode inte-

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Fig. 19. This exploded view of the dialysis probe connector shows the double lumen probe with the perfusion liquid flow passing via the outer capillary into the membrane region and finally flowing through the inner capillary into the flowthrough cell with integrated chloride sensor [111] (reprinted from [111] with permission. Copyright 2000 Elsevier Science).

grated into a flow-cell in the silicon part of the device, which consists of two polymer-bonded silicon wafers anisotropically etched in KOH. In comparison to a conventional flow-through sensor for chloride, whose response time was 60 s at a flow rate of 1 ml s − 1, the miniaturized version is much faster (11 s at the same flow rate). This is largely due to the low dead volume of the system. A microfluidic device integrating a dialysis membrane appeared recently in the literature, developed for desalting of oligonucleotide samples for electrospray ionization mass spectrometry (ESI-MS) [112,113]. The advantage of miniaturization in this case lies in the reduced flow rates possible while still maintaining a high desalting efficiency, which improves both dialysis time and MS sensitivity. The device concept, shown in Fig. 20c, is straightforward, using polymer substrates structured by laser ablation or laser cutting. The chip consists of a top and bottom plate, both containing a serpentine channel laid out in such a

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way that the two channels mate perfectly when the plates are assembled. A piece of dialysis membrane is sandwiched between the two aligned planes when the chip is sealed. The sample channel is 160 mm wide while the perfusion channel measures 500 mm, thereby increasing the diffusional mass transport across the membrane. For operation, the sample and perfusate streams flow in opposite directions. In this way, the cleanest perfusate comes in contact with already-somewhat-purified sample, ensuring a larger concentration gradient and stronger diffusion as a result. Fig. 20a and b shows a 40-fold improvement of the signal-to-noise ratio of an ESI-mass spectrum of a 5 mM horse heart myoglobin sample obtained by on-line dialysis in the microfabricated device.

Fig. 20. (A) ESI-mass spectrum of 5 mM horse heart myoglobin in 500 mM NaCl, 100 mM Tris, and 10 mM EDTA, from direct infusion, compromised by sodium adduction. (B) Same sample after on-line dialysis in the microchip using 10 mM NH4OAc and 1% acetic acid as countercurrent buffer: the signal-to-noise ratio improves more than 40-fold. (C) Photo and schematic cross-section of the dialysis chip with a sample channel on top, an intermediate dialysis membrane and a buffer channel in the lower part [112] (reprinted from [112] with permission. Copyright 1998 American Chemical Society).

3.4.4. Liquid–liquid extraction Only a few examples for integrated liquid–liquid extraction (LLE) systems exist, although miniaturized systems offer the advantages of operation in the laminar flow regime and the possibility of creating large interfacial contact regions relative to the small volumes being manipulated. Macroscopic LLE systems mostly rely on turbulent mixing and are thus prone to emulsion or foam generation. Miniaturized LLE devices have been fabricated by anodic bonding of silicon and glass substrates with etched, matching channels in a meander layout [114]. A slight horizontal misalignment between the two 100 mm wide channels was introduced on purpose to limit the width of the contact area between the two streams to around 10 mm. LLE units were fabricated in arrays to increase the throughput, which was in the order of 1 ml min − 1. The device was evaluated for a range of extraction processes from organic to aqueous solutions, for targets including substituted nitrobenzoic acid, tocopherol succinate, and arginine–cyclohexylamine salt. Very recently, a multiphase microreactor based on a simple glass device consisting of a cross layout of microchannels 380 mm wide and 380 mm deep, formed using a slitting saw rather than by etching, was reported [115]. In this case, the researchers opted for the use of slug flow of two immiscible liquids rather than side-by-side laminar flows to carry out extraction of material from

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one phase to the other. This is because internal circulatory flow is generated in the slugs as they pass along the channel, creating a large enhancement in interfacial mass transfer and hence reaction rate. The utility of the concept was demonstrated by monitoring the extraction of acetic acid from kerosene slugs into aqueous slugs containing KOH and a pH indicator. An ion-pair solvent extraction was performed by Tokeshi et al. [116] to extract Fe(II) from an aqueous solution of Fe-bathophenanthrolinedisulfonic acid complex into a chloroform solution of tri-n-octylmethylammonium chloride. Both solutions were introduced via separate channels on a quartz chip, merged to develop a two-phase laminar flow and separated after 10 mm. The extraction time in a 250 mm channel was 45 s, which is at least 1 order of magnitude shorter than conventional extraction methods.

3.4.5. Free-flow electrophoresis Free-flow electrophoresis (FFE) is an electrophoretic separation technique that relies on an electric field perpendicular to the overall flow direction in the system. In the conventional instrument, the sample is introduced in a continuous fashion at one point along the top of the two-dimensional separation bed and carried through the bed by pressure-driven flow. Membranes along the sides of the bed separate it from two chambers, one on either side, which run adjacent to the bed along its entire length. Electrodes placed in these side chambers are used to apply an electric field across the bed perpendicular to the direction of flow. The result is deflection of charged species from the direction of flow, as shown in Fig. 21a, with the angle of deflection increasing as electrophoretic mobility and/or electric field strength increases. At the end of the bed, the eluting streams of individual components may be collected for further analysis. FFE was originally developed for the preparative scale separation and isolation of cells, bacteria, viruses, proteins and other biological entities [117]. The continuous nature of the separation, and the capability of isolating sample components, make this an attractive ‘smart filter’ for lab-on-chip systems as well.

Fig. 21. (a) Principle of free-flow electrophoresis. The sample is introduced continuously at the inlet, carried through the separation bed by pressure-driven flow and deflected, depending on the charge-to-mass ratio, by a perpendicular electric field. (b) Inlet section of an FFE chip. Arrays of V-grooves at the sides act as a ‘membrane’ to apply the separation voltage, while at the inlet side they assure undisturbed flow of the sample and separation buffer [118] (reprinted from [118] with permission. Copyright 1994 American Chemical Society).

A silicon-based miniaturized FFE system with a 10 mm wide, 50 mm long and 50 mm deep separation bed (25 ml volume) was reported by Raymond et al. in 1994 [118]. The fluidic structure was fabricated by anisotropic wet etching and sealed with a glass plate using anodic bonding. A

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three-dimensional view of the inlet region is shown in Fig. 21b. Both in- and outlets are formed as arrays of V-groove channels to ensure an even distribution of solution over the width of the bed. A few channels in the inlet array are used to continuously introduce a sample stream into the device, while the others carry the buffer electrolyte. In this microfabricated embodiment of FFE, the membranes isolating the electrode chambers from the bed are mimicked by arrays of 2500 shallow grooves along the length of the bed. Proof-of-concept was demonstrated by continuously separating a mixture of three fluorescently labeled amino acids, with a residence time of 73 s at an applied voltage of 50 V and a constant flow rate of 50 ml min − 1. The separation of high molecular weight compounds is of special interest for sample pretreatment applications. At a low field strength of 25 V cm − 1, human serum albumin (MW 69526) could be isolated from bradykinin (MW 1436) and ribonuclease A (MW 14453). More complex samples like tryptic digests could also be analyzed [119]. In a slightly modified version of the chip with additional outlet holes, a fraction collection experiment with whole rat plasma was performed. More recently, an FFE device formed from two Plexiglas plates separated by a 100 mm spacer was described for continuous on-line separation in conjunction with flow injection analysis and chromatographic systems [120]. Using fluorescently labeled biotin as a model ligand and streptavidin as affinity protein, it was possible to separate the labeled affinity complex from the free ligand in under 2 min. Fluorescence detection was facilitated by the optical transparency of the device. The applicability of the FFE device in combination with an HPLC separation was also demonstrated, again in the analysis of biotin in human urine at micromolar levels.

3.4.6. Cell lysis and purification An ultimate goal for microanalytical device development is the analysis of whole blood or integral cells on-chip. However, this requires the analyte of interest to be freed from the surrounding cell membrane and related structural material by lysis or disruption. Four approaches for on-

chip lysis are predominant, namely chemical lysis using a detergent, thermal lysis, ultrasonication to physically break open spores, and electroporation. Li and Harrison reported in 1997 on the possibility of transporting, manipulating and modifying biological cells on planar glass substrates using EOF and electrophoretic flow [121]. Cell lysis was demonstrated by mixing a sample stream containing canine erythocytes in an isotonic buffer with a 3 mM sodium dodecyl sulfate (SDS) detergent solution at a T-intersection. Efficient cell lysis could be observed by the absence of light scattering by intact cells shortly after the mixing and by video microscopy. Other options for chemical lysis were less suitable; the addition of copper(II) ions because of the slow reaction kinetics, and acidic solutions because of the EOF reversal associated with low pH. A further step in integration is the combination of cell lysis with multiplex PCR and electrophoretic sizing published by Waters et al. [78]. As PCR requires thermocycling anyway, a 4 min heat treatment of the Escherichia coli sample at 94 °C was selected as a combined lysis and denaturation step for the first PCR cycle. Both lysis and amplification were carried out on-chip in a commercial thermal cycler. Lysis of targets like bacterial spores is more challenging, as the spore outer cortex is very resistant to both chemical and physical treatment. Belgrader et al. [122] therefore selected a harsh physical force to break the spores open. Ultrasonic waves (power 40 W, frequency 47 kHz) were coupled into the microfluidic system containing the sample in a suspension with 106 mm diameter glass beads. An external ultrasonic transducer based on a stack of piezoelectric transducer disks was mechanically interfaced via a flexible membrane to the polymer chip using a titanium horn to transfer the ultrasonic energy into the lysis chamber. It was proposed that gaseous cavitation by collapsing bubbles from dissolved gas creates localized high pressure waves that disrupt the spores. The efficiency of the disruption was verified by subsequent on-chip, real-time PCR of the lysis product of a 105 Bacillus subtilis spores ml − 1 sample. TaqMan analysis was used, and the threshold cycle, CT (the point at which a signifi-

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cant increase in positive signal is suddenly observed), was determined. Purified spores ultrasonically disrupted for 30 s yielded a CT of 28 (14 min), while the CT for amplification of untreated spores appeared only after 40 cycles (21 min). The efficiency of the miniaturized sonicator was nearly identical to a conventional 1 h germination procedure used in labs today. An improved, self-contained version of the cartridge developed for portable applications has also been presented [123]. Further optimization of the ultrasonic energy transfer via a flexible membrane into the chamber by applying a static pressure to the sample-bead suspension in the lysis chamber reduced the CT further down to 24 [124]. In electroporation, cell membranes are reversibly or irreversibly rendered permeable by applying a high electric field across the cell. This technique has been applied on-chip to study cell permeabilization [125] and to introduce plasmid DNA from yeast to E. coli cells [126].

4. Conclusion It is clear that the integration of sample pretreatment into microfluidic devices represents one of the remaining hurdles towards achieving true mTAS. The challenge is made more complex by the enormous variation in samples to be analyzed. Moreover, the pretreatment technique has to be compatible with the analysis device to which it is coupled in terms of time, reagent and power consumption as well as sample volume. As this review shows, however, significant progress has been made, especially in recent years, driven largely by developments in the life sciences. An exponential increase in papers dealing with this subject is to be expected in the next couple of years.

References [1] H.M. Widmer, Trends Anal. Chem. 2 (1983) VIII – X. [2] A. Manz, N. Graber, H.M. Widmer, Sens. Actuators B 1 (1990) 244 – 248.

263

[3] A. Manz, D.J. Harrison, E.M.J. Verpoorte, J.C. Fettinger, A. Paulus, H. Lu¨ di, H.M. Widmer, J. Chromatogr. 593 (1992) 253 – 258. [4] D.J. Harrison, K. Fluri, K. Seiler, Z.H. Fan, C.S. Effenhauser, A. Manz, Science 261 (1993) 895 – 897. [5] S.C. Jacobson, R. Hergenro¨ der, L.B. Koutny, J.M. Ramsey, Anal. Chem. 66 (1994) 1114 – 1118. [6] A. Manz, C.S. Effenhauser, N. Burggraf, D.J. Harrison, K. Seiler, K. Fluri, J. Micromech. Microeng. 4 (1994) 257 – 265. [7] G.J.M. Bruin, Electrophoresis 21 (2000) 3931 – 3951. [8] R.P. Manginell, G.C. Frye-Mason, R.J. Kottenstette, P.R. Lewis, C.C. Wong, Proceedings of the Solid-State Sensor and Actuator Workshop, Hilton Head Island, SC, 2000, pp. 179 – 182. [9] C.S. Effenhauser, A. Manz, H.M. Widmer, Anal. Chem. 65 (1993) 2637 – 2642. [10] S.C. Jacobson, L.B. Koutny, R. Hergenro¨ der, A.W. Moore, J.M. Ramsey, Anal. Chem. 66 (1994) 3472 – 3476. [11] D.J. Harrison, Z. Fan, K. Fluri, K. Seiler, Proceedings of the IEEE Solid-State Sensor and Actuator Workshop, Hilton Head Island, SC, IEEE Electron Devices Society, 1994, pp. 21 – 24. [12] K. Fluri, G. Fitzpatrick, N. Chiem, D.J. Harrison, Anal. Chem. 68 (1996) 4285 – 4290. [13] K. Seiler, Z.H. Fan, K. Fluri, D.J. Harrison, Anal. Chem. 66 (1994) 3485 – 3491. [14] S.C. Jacobson, R. Hergenro¨ der, A.W. Moore, J.M. Ramsey, Anal. Chem. 66 (1994) 4127 – 4132. [15] J.C.T. Eijkel, A. Prak, S. Cowen, D.H. Craston, A. Manz, J. Chromatogr. 815 (1998) 265 – 271. [16] J.P. Kutter, R.S. Ramsey, S.C. Jacobson, J.M. Ramsey, J. Microcolumn Sep. 10 (1998) 313 – 319. [17] D.F. Swaile, M.J. Sepaniak, Anal. Chem. 63 (1991) 179 – 184. [18] Y.J. Liu, R.S. Foote, S.C. Jacobson, R.S. Ramsey, J.M. Ramsey, Anal. Chem. 72 (2000) 4608 – 4613. [19] S.D. Mangru, D.J. Harrison, Electrophoresis 19 (1998) 2301 – 2307. [20] E.M.J. Verpoorte, B.H. van der Schoot, S. Jeanneret, A. Manz, N.F. de Rooij, Interfacial Des. Chem. Sensing 561 (1994) 244 – 254. [21] E.M.J. Verpoorte, B.H. van der Schoot, S. Jeanneret, A. Manz, H.M. Widmer, N.F. de Rooij, J. Micromech. Microeng. 4 (1994) 246 – 256. [22] A. Daridon, M. Sequeira, G. Pennarun-Thomas, H. Dirac, J.P. Krog, P. Gravesen, J. Lichtenberg, D. Diamond, E. Verpoorte, N.F. de Rooij, Sens. Actuators B 76 (2001) 235 – 243. [23] A. Daridon, V. Fascio, J. Lichtenberg, R. Wu¨ trich, H. Langen, E. Verpoorte, N.F de Rooij, Fresenius J. Anal. Chem. 371 (2001) 261 – 269. [24] G.N. Doku, S.J. Haswell, Anal. Chim. Acta 382 (1999) 1 – 13. [25] S.J. Haswell, Analyst 122 (1997) R1 – R10.

264

J. Lichtenberg et al. / Talanta 56 (2002) 233–266

[26] F.E.P. Mikkers, F.M. Everearts, T.P.E.M. Verheggen, J. Chromatogr. 169 (1979) 11 –20. [27] D.S. Burgi, R.-L. Chien, J. Microcolumn Sep. 3 (1991) 199 –202. [28] R.-L. Chien, D.S. Burgi, Anal. Chem. 64 (1992) 489A – 496A. [29] D.S. Burgi, R.-L. Chien, Anal. Chem. 63 (1991) 2042 – 2047. [30] R.-L. Chien, J.C. Helmer, Anal. Chem. 63 (1991) 1354 – 1361. [31] J. Lichtenberg, E. Verpoorte, N.F. de Rooij, Electrophoresis 22 (2001) 258 –271. [32] J. Li, C. Wang, J.F. Kelly, D.J. Harrison, P. Thibault, Electrophoresis 21 (2000) 198 –210. [33] A.E. Herr, J.I. Molho, J.G. Santiago, M.G. Mungal, T.W. Kenny, Anal. Chem. 72 (2000) 1053 –1057. [34] S.C. Jacobson, J.M. Ramsey, Electrophoresis 16 (1995) 481 – 486. [35] S.C. Jacobson, R. Hergenro¨ der, L.B. Koutny, R.J. Warmack, M. Ramsey, Anal. Chem. 66 (1994) 1107 – 1113. [36] H. Yang, R.-L. Chien, J. Chromatogr. A 924 (2001) 155 – 163. [37] J.P. Quirino, S. Terabe, Science 282 (1998) 465 – 468. [38] J.P. Quirino, S. Terabe, Anal. Chem. 72 (2000) 1023 – 1030. [39] J. Palmer, N.J. Munro, J.P. Landers, Anal. Chem. 71 (1999) 1679 – 1687. [40] J. Palmer, D.S. Burgi, J.P. Landers, Anal. Chem. 73 (2001) 725 – 731. [41] F.M. Everaerts, J.L. Beckers, T.P.E.M. Verheggen, Isotachophoresis: Theory, Instrumentation, and Applications, Elsevier, New York, 1976. [42] R.A. Mosher, D.A. Saville, W. Thormann, The Dynamics of Electrophoresis, VCH, Weinheim, 1992. [43] D. Kaniansky, M. Masa´ r, J. Bielikova´ , F. Iva´ nyi, F. Eisenbeiss, B. Stanislawski, B. Grass, A. Neyer, M. Jo¨ hnck, Anal. Chem. 72 (2000) 3596 –3604. [44] B. Graß, A. Neyer, M. Jo¨ hnck, D. Siepe, F. Eisenbeiß, G. Weber, R. Hergenro¨ der, Sens. Actuators B 72 (2001) 249 –258. [45] J.E. Prest, S.J. Baldock, P.R. Fielden, B.J.T. Brown, Analyst 126 (2001) 433 –437. [46] J.P. Kutter, S.C. Jacobson, J.M. Ramsey, J. Microcolumn Sep. 12 (2000) 93 –97. [47] J.P. Kutter, S.C. Jacobson, N. Matsubara, J.M. Ramsey, Anal. Chem. 70 (1998) 3291 –3297. [48] R.D. Oleschuk, L.L. Shultz-Lockyear, Y.B. Ning, D.J. Harrison, Anal. Chem. 72 (2000) 585 –590. [49] R.D. Oleschuk, A.B. Jemere, L.L. Schultz-Lockyear, F. Fajuyigbe, D.J. Harrison, Proceedings of the Micro Total Analysis Systems Symposium 2000, Kluwer Academic, Dordrecht, 2000, pp. 11 – 14. [50] D. Figeys, R. Aebersold, Anal. Chem. 70 (1998) 3721 – 3727. [51] R. Boom, C.J.A. Sol, M.M.M. Salimans, C.L. Jansen, P.M.E. Wertheim-van Dillen, J. van der Noordaa, J. Clin. Microbiol. 28 (1990) 495 –503.

[52] L.A. Christel, K. Petersen, W. McMillan, M.A. Northrup, J. Biomech. Eng. 121 (1999) 22 – 27. [53] J. Khandurina, S.C. Jacobson, L.C. Waters, R.S. Foote, J.M. Ramsey, Anal. Chem. 71 (1999) 1815 – 1819. [54] J. Khandurina, T.E. McKnight, S.C. Jacobson, L.C. Waters, R.S. Foote, J.M. Ramsey, Anal. Chem. 72 (2000) 2995 – 3000. [55] Y.-C. Lin, H.-C. Ho, C.-K. Tseng, S.-Q. Hou, J. Micromech. Microeng. 11 (2001) 189 – 194. [56] G. Jiang, D.J. Harrison, Analyst 125 (2000) 2176 – 2179. [57] Z.H. Fan, S. Mangru, R. Granzow, P. Heaney, W. Ho, Q.P. Dong, R. Kumar, Anal. Chem. 71 (1999) 4851 – 4859. [58] J.-W. Choi, C.H. Ahn, S. Bhansali, H.T. Henderson, Sens. Actuators B 68 (2000) 34 – 39. [59] A. Dodge, K. Fluri, E. Verpoorte, N.F. de Rooij, Anal. Chem. 73 (2001) 3400 – 3409. [60] K. Sato, M. Tokeshi, T. Odake, H. Kimura, T. Ooi, M. Nakao, T. Kitamori, Anal. Chem. 72 (2000) 1144 – 1147. [61] K. Sato, M. Tokeshi, H. Kimura, T. Kitamori, Anal. Chem. 73 (2001) 1213 – 1218. [62] K. Mullis, F. Faloona, S. Scharf, R. Saiki, G. Horn, H. Erlich, Cold Spring Harbor Symp. Quant. Biol. 51 (1986) 263 – 273. [63] M.A. Northrup, M.T. Ching, R.M. White, R.T. Watson, Proceedings of the Transducers’93: Seventh International Conference on Solid State Sensors and Actuators, Yokohama, Japan, 1993, pp. 924 – 927. [64] P. Wilding, M.A. Shoffner, L.J. Kricka, Clin. Chem. 40 (1994) 1815 – 1818. [65] M.A. Shoffner, J. Cheng, G.E. Hvichia, L.J. Kricka, P. Wilding, Nucleic Acids Res. 24 (1996) 375 – 379. [66] B.C. Giordano, E.R. Copeland, J.P. Landers, Electrophoresis 22 (2001) 334 – 340. [67] T.B. Taylor, E.S. Winn-Deen, E. Picozza, T.M. Woudenberg, M. Albin, Nucleic Acids Res. 25 (1997) 3164 – 3168. [68] M.A. Northrup, B. Benett, D. Hadley, P. Landre, S. Lehew, J. Richards, P. Stratton, Anal. Chem. 70 (1998) 918 – 922. [69] P. Belgrader, S. Young, B. Yuan, M. Primeau, L.A. Christel, F. Pourahmadi, A. Northrup, Anal. Chem. 73 (2001) 286 – 289. [70] T.M. Lee, I.M. Hsing, A.I. Lao, M.C. Carles, Anal. Chem. 72 (2000) 4242 – 4247. [71] M.A. Burns, C.H. Mastrangelo, T.S. Sammarco, F.P. Man, J.R. Webster, B.N. Johnson, B. Foerster, D. Jones, Y. Fields, A.R. Kaiser, D.T. Burke, Proc. Natl Acad. Sci. USA 93 (1996) 5556 – 5561. [72] M.A. Burns, V.N. Johnson, S.N. Brahmasandra, K. Handique, J.R. Webster, M. Krishnan, T.S. Sammarco, P.M. Man, D. Jones, D. Heldsinger, C.H. Mastrangelo, D.T. Burke, Science 282 (1998) 484 – 487. [73] R.P. Oda, M.A. Strausbauch, A.F.R. Huhmer, N. Borson, S.R. Jurrens, J. Craighead, P.J. Wettstein, B. Eckloff, B. Kline, J.P. Landers, Anal. Chem. 70 (1998) 4361 – 4368.

J. Lichtenberg et al. / Talanta 56 (2002) 233–266 [74] B.C. Giordano, J. Ferrance, S. Swedberg, A.F. Huhmer, J.P. Landers, Anal. Biochem. 291 (2001) 124 – 132. [75] A.T. Woolley, D. Hadley, P. Landre, A.J. de Mello, R.A. Mathies, M.A. Northrup, Anal. Chem. 68 (1996) 4081 – 4086. [76] E.T. Lagally, P.C. Simpson, R.A. Mathies, Sens. Actuators B 63 (2000) 138 –146. [77] E.T. Lagally, I. Medintz, R.A. Mathies, Anal. Chem. 73 (2001) 565 – 570. [78] L.C. Waters, S.C. Jacobson, N. Kroutchinina, J. Khandurina, R.S. Foote, J.M. Ramsey, Anal. Chem. 70 (1998) 158 – 162. [79] L.C. Waters, S.C. Jacobson, N. Kroutchinina, J. Khandurina, R.S. Foote, J.M. Ramsey, Anal. Chem. 70 (1998) 5172 – 5176. [80] J.W. Hong, T. Fujii, M. Seki, T. Yamamoto, I. Endo, Electrophoresis 22 (2001) 328 –333. [81] M.U. Kopp, A.J. de Mello, A. Manz, Science 280 (1998) 1046 – 1048. [82] S.C. Jacobson, J.M. Ramsey, Anal. Chem. 68 (1996) 720 – 723. [83] N. Gottschlich, C.T. Culbertson, T.E. McKnight, S.C. Jacobson, M. Ramsey, J. Chromatogr. B 745 (2000) 243 – 249. [84] Q. Xue, Y.M. Dunnayevskiy, F. Foret, B.L. Karger, Rapid Commun. Mass Spectrom. 11 (1997) 1253 – 1256. [85] C. Wang, R. Oleschuk, F. Ouchen, J.J. Li, P. Thibault, D.J. Harrison, Rapid Commun. Mass Spectrom. 14 (2000) 1377 – 1383. [86] M.J. Heller, E. Tu, A.H. Forster, Electrophoresis 21 (2000) 157 – 164. [87] R. Radtkey, L. Feng, M. Muralhider, M. Duhon, D. Canter, D. DiPierro, S. Fallon, E. Tu, K. McElfresh, M. Nerenberg, R. Sosnowski, Nucleic Acids Res. 28 (2000) e17. [88] L. Westin, X. Xu, C. Miller, L. Wang, C.F. Edman, M. Nerenberg, Nature Biotechnol. 18 (2000) 199 –204. [89] P. Wilding, L.J. Kricka, J. Cheng, G. Hvichia, M.A. Shoffner, P. Fortina, Anal. Biochem. 257 (1998) 95 – 100. [90] H. Andersson, W.v.d. Wijngaart, P. Enoksson, G. Stemme, Sens. Actuators A 67 (2000) 203 –208. [91] L. Ceriotti, E. Verpoorte, N.F. de Rooij, Anal. Chem. (2002). [92] G. Kittilsland, G. Stemme, Sens. Actuators A 21 –23 (1990) 904 – 907. [93] W.-H. Chu, R. Chin, T. Huen, M. Ferrari, J. Microelectromech. Syst. 8 (1999) 34 – 42. [94] J.K. Tu, T. Huen, R. Szema, M. Ferrari, Biomed. Microdevices 1 (1999) 113 –120. [95] X. Xing, J.M. Yang, Y.C. Tai, C.M. Ho, Sens. Actuators A 73 (1999) 184 –191. [96] H. Seidel, J. Electrochem. Soc. 137 (1990) 3626 –3632. [97] S. Kuiper, H.v. Wolferen, C.v. Rijn, W. Nijdam, G. Krijnen, M. Elwenspoek, J. Micromech. Microeng. 11

265

(2001) 33 – 37. [98] B. He, L. Tan, F. Regnier, Anal. Chem. 71 (1999) 1464 – 1468. [99] G.T.A. Kovacs, Micromachined Transducers Sourcebook, McGraw-Hill, New York, 1998. [100] P. Gravesen, J. Branebjerg, O.S. Jenson, J. Micromech. Microeng. 3 (1993) 168 – 182. [101] B.H. Weigl, M.R. Holl, D. Schutte, J.P. Brody, P. Yager, Proceedings of the Second International Symposium on Miniaturized Total Analysis Systems mTAS96, AMI, Basel, 1996, pp. 174 – 184. [102] J.P. Brody, T.D. Osborn, F.K. Foster, P. Yager, Sens. Actuators A 54 (1996) 704 – 708. [103] J.P. Brody, P. Yager, Sens. Actuators A 58 (1997) 13 – 18. [104] B.H. Weigl, P. Yager, Science 283 (1999) 346 – 347. [105] A.E. Kamholz, B.H. Weigl, B.A. Finlayson, P. Yager, Anal. Chem. 71 (1999) 5340 – 5347. [106] C.E. Lunte, D.O. Scott, P.T. Kissinger, Anal. Chem. 63 (1991) 773A – 779A. [107] U. Ungerstedt, J. Inter. Med. 230 (1991) 365 – 373. [108] N. Toro, T. Laurell, L. Gorton, G. Marko-Varga, Anal. Chim. Acta 379 (1999) 281 – 305. [109] R. Freaney, A. McShane, T.V. Keaveny, M. McKenna, K. Rabenstein, F.W. Scheller, D. Pfeiffer, G. Urban, I. Moser, G. Jobst, A. Manz, E. Verpoorte, M.W. Widmer, D. Diamond, E. Dempsey, F.J. Saez de Viteri, M. Smyth, Ann. Clin. Biochem. 34 (1997) 291 – 302. [110] E. Dempsey, D. Diamond, M. Smyth, G. Urban, G. Jobst, I. Moser, E.M.J. Verpoorte, A. Manz, H.M. Widmer, K. Rabenstein, R. Freaney, Anal. Chim. Acta 346 (1997) 341 –349. [111] S. Bo¨ hm, W. Olthuis, P. Bergveld, Sens. Actuators B 63 (2000) 201 – 208. [112] N.X. Xu, Y.H. Lin, S.A. Hofstadler, D. Matson, C.J. Call, R.D. Smith, Anal. Chem. 70 (1998) 3553 – 3556. [113] F. Xiang, Y.H. Lin, J. Wen, D.W. Matson, R.D. Smith, Anal. Chem. 71 (1999) 1485 – 1490. [114] J. Shaw, R. Nudd, B. Naik, C. Turner, D. Rudge, M. Benson, A. Garman, Proceedings of the Micro Total Analysis Systems 2000 Symposium, Enschede, Kluwer Academic, Dordrecht, 2000, pp. 371 – 374. [115] J.R. Burns, C. Ramshaw, Lab Chip 1 (2001) 10 – 15. [116] M. Tokeshi, T. Minagawa, T. Kitamori, Anal. Chem. 72 (2000) 1711 – 1714. [117] K. Hanning, H.G. Heidrich, Free-flow electrophoresis, GIT Verlag GmbH, Darmstadt, Germany, 1990. [118] D.E. Raymond, A. Manz, H.M. Widmer, Anal. Chem. 66 (1994) 2858 – 2865. [119] D.E. Raymond, A. Manz, H.M. Widmer, Anal. Chem. 68 (1996) 2515 – 2522. [120] M. Mazereeuw, C.M. de Best, U.R. Tjaden, H. Irth, J. van der Greef, Anal. Chem. 72 (2000) 3881 – 3886. [121] P.C.H. Li, D.J. Harrison, Anal. Chem. 69 (1997) 1564 – 1568. [122] P. Belgrader, D. Hansford, G.T.A. Kovacs, K.

266

J. Lichtenberg et al. / Talanta 56 (2002) 233–266

Venkateswaran, R. Mariella, F. Milanovich, S. Nasarabadi, M. Okuzumi, F. Pourahmadi, M.A. Northrup, Anal. Chem. 71 (1999) 4232 –4236. [123] P. Belgrader, M. Okuzumi, D.A. Borkholder, F. Pourahmadi, M.A. Northrup, Biosens. Bioelectron. 12 (2000) 849 – 852. [124] M.T. Taylor, P. Belgrader, B.J. Furman, F. Pourah-

madi, G.T.A. Kovacs, M.A. Northrup, Anal. Chem. 73 (2001) 492 – 496. [125] Y. Huang, B. Rubinsky, Sens. Actuators A 89 (2001) 242 – 249. [126] R. Poppe, Proceedings of the International Conference on Microreaction Technology (IMRET) 3, Frankfurt/ M., Springer, 1999, pp. 461 – 468.