Microfluidics meets MEMS - Proceedings of the IEEE

Microfluidics Meets MEMS
ELISABETH VERPOORTE AND NICO F. DE ROOIJ, FELLOW, IEEE
Invited Paper
The use of planar fluidic devices for performing small-volume
chemistry was first proposed by analytical chemists, who coined
the term “miniaturized total chemical analysis systems” (TAS) for
this concept. More recently, the TAS field has begun to encompass other areas of chemistry and biology. To reflect this expanded
scope, the broader terms “microfluidics” and “lab-on-a-chip” are
now often used in addition to TAS. Most microfluidics researchers
rely on micromachining technologies at least to some extent to produce microflow systems based on interconnected micrometer-dimensioned channels. As members of the microelectromechanical
systems (MEMS) community know, however, one can do more with
these techniques. It is possible to impart higher levels of functionality by making features in different materials and at different levels
within a microfluidic device. Increasingly, researchers have considered how to integrate electrical or electrochemical function into
chips for purposes as diverse as heating, temperature sensing, electrochemical detection, and pumping. MEMS processes applied to
new materials have also resulted in new approaches for fabrication
of microchannels. This review paper explores these and other developments that have emerged from the increasing interaction between
the MEMS and microfluidics worlds.
Keywords—Lab-on-a-chip, microfluidics, microfabrication,
micromachining, miniaturized total chemical analysis systems
(TAS), review .
I. INTRODUCTION
Though there exist early, pre-1990 examples of microelectromechanical systems (MEMS)-type microfluidic devices
for chemical applications, it was the groundbreaking paper
of Manz et al. [1] in 1990 that established the field of miniaturized total chemical analysis systems ( TAS). The TAS
concept is an extension of the total chemical analysis system
concept (TAS), which was put forth in the early 1980s to
address the issue of automation in analytical chemistry [2].
Manuscript received February 26, 2003; revised April 4, 2003.
E. Verpoorte was with the Institute of Microtechnology, University of
Neuchâtel, CH-2007 Neuchâtel, Switzerland. She is now at the Groningen
Research Institute of Pharmacy, University of Groningen, 9713 AV
Groningen, The Netherlands (e-mail: [email protected]).
N. F. de Rooij is with the Institute of Microtechnology, University of Neuchâtel, CH-2007 Neuchâtel, Switzerland (e-mail:
[email protected]).
Digital Object Identifier 10.1109/JPROC.2003.813570
To simplify the job of the analytical chemist, the TAS concept proposed the full incorporation of analytical procedures
into flowing systems. Carrier streams of fluids, rather than
human hands, take over the role of sample transport between
different sample manipulation steps. Flow paths are defined
by interconnected pieces of tubing, arranged in such a way
that the desired analysis can be performed. The TAS is a
smaller, faster version of this, with analyses taking place
in flow systems having L and even sub- L volumes, to
achieve times on the order of seconds rather than many minutes. The devices used are small and monolithic in nature.
Three-dimensional (3-D) tubing-based systems are replaced
by networks of microchannels integrated into planar substrate surfaces, with cross-sectional dimensions generally
on the order of micrometers. Though still in its infancy, the
interest in this technology has grown explosively over the
last decade, with an increasing number of special symposia
and conferences dedicated to the subject. Researchers from
many disciplines other than analytical chemistry have also
wholeheartedly embraced the fundamental fluidic principle
of TAS as a way of developing new research tools for
chemical and biological applications. This has led, among
other things, to the emergence of TAS-like fluidic structures in microreactor technology. It has also resulted in the
introduction of new terminologies and concepts. “Microfluidics,” or alternatively, “lab-on-a-chip” (LOC), are recent
monikers that have entered the TAS jargon, and are increasingly used in place of TAS as general terms for the
field. This development is in fact a tacit acknowledgment,
perhaps, that the use of microflow systems has long since
ceased to be the domain of the analytical chemist alone.
The ability to make networks of interconnecting channels
is what lies at the heart of microfluidics, and makes it so
powerful. The crucial enabling technologies came first from
the world of MEMS, where the use of photolithographic
processes to obtain micrometer features in silicon and other
substrates was well established. The first successful demonstration of chip-based analysis in the early nineties involved
the fast separation of fluorescent dyes [3], [4] and fluorescently labeled amino acids [5] by capillary electrophoresis
(CE). This technique, based on the separation of charged
0018-9219/03$17.00 © 2003 IEEE
930
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 1. Video imaging of sample loading, injection, and separation
by micellar electrokinetic capillary chromatography (MECC) in a
simple cross injector geometry. Separation is effected by interaction
of the analyte with micelles composed of the surfactant, sodium
dodecyl sulfate (SDS), which was added to the run buffer. With
their hydrophobic interiors and charged surfaces, the micelles
are aggregates which act as a pseudostationary phase transported
electrokinetically in the electric field. Channels are 40 m wide
and 10 m deep, so that the injected plug volume is 20 pL. The
vertical channel is filled with sample containing 2 mM each
of glycine (Gly) and arginine (Arg), both fluorescently labeled
with fluorescein isothiocyanate (FITC). The horizontal channel
contains MECC buffer [75-mM SDS, 10-mM Na HPO , and
6-mM Na B O (pH 9.2)]. Arrows indicate direction and relative
magnitudes of the flows induced by application of voltages at the
ends of these channels. The spot visible in the intersection in images
C and D is due to reflection of laser light from an imperfection in
the microchannel surface. (Reprinted with permission from [7].
Copyright 1996 American Chemical Society.)
species according to their different mobilities in an applied
electric field, is performed in liquid-filled capillaries. CE
is particularly well suited to the chip format for a number
of reasons. For one thing, the electric fields used to separate species electrophoretically also generate a bulk flow of
liquid, known as electroosmosis [6]. Electroosmotic flow
(EOF) is thus the mechanism by which liquids are moved
from one end of the separation capillary to the other, obviating the need for mechanical pumps and valves. This
makes this technique very amenable to miniaturization, as
it is far simpler to make an electrical contact to a chip via
a wire immersed in a reservoir than it is to make a robust
connection to a pump. More important, however, is that all
the basic fluidic manipulations that a chemist requires for
microchip electrophoresis, or any other liquid handling for
that matter, have been adapted to electrokinetic microfluidic
chips. This is illustrated in Fig. 1, which shows the formation of a 20–pL injection plug containing two fluorescent
species and the subsequent separation of the two analytes
in just 160 ms [7].
Precise liquid pumping and handling may also be accomplished using applied pressure or centrifugal force. As an
illustration, Fig. 2 shows the squeezing or “hydrodynamic
focusing” of a fluorescent sample stream at a channel intersection using two buffer flows from side channels [8]. Pressures were applied at the ends of the inlet and side channels
to generate liquid flows. The width of the resulting focused
sample stream is determined by the ratio of pressures applied
to the side channel and to the inlet. Widths as small as 50 nm
were observed, ensuring fast diffusion of molecules in the
side streams across the sample stream to achieve ultrafast
s mixing. It is also clear that controlled s dilution may
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
Fig. 2. (a) Fluorescence microscope image of hydrodynamic
focusing in an intersection of microchannels made in silicon (the
channels are outlined with a dashed line). The side and outlet
channels are 10 m wide, while the inlet channel terminates in a
2-m nozzle. All channels are 10 m deep. The sample contained
5-carboxyfluorescein, a fluorescent dye. w is the width of the
channel; w is the width of the focused inlet stream. Inlet pressure
is 5 psi; side pressure is 5.5 psi. (b) Images of the 3-D structure of
the focused sample stream as a function of side pressure, obtained
by confocal scanning microscopy. Inlet pressure is 5 psi. Side
pressure of (a) is 2.5 psi; (b), 5.0 psi; (c), 5.5 psi; and (d), 6.0 psi.
(Reprinted with permission from [8]. Copyright 1998 American
Physical Society.)
be carried out in this fashion. As this example proves, pressure-driven flow may also be effectively used to carry out
the necessary liquid handling required for chemical applications, though it is somewhat more difficult to implement in
conjunction with microfluidic devices. The use of capillary
forces or well-defined surface tension gradients is also attracting more attention, as this is a passive means of moving
liquids in small channels.
One of the most evident trends in the microfluidics world
is that of increasing complexity in the microchannel architectures being used. Since the majority of devices do not feature on-board valves, this requires very precise control in all
branches of the networks, a very challenging aspect of this
technology. Some novel valving approaches have been proposed, which may facilitate development of these systems.
The true TAS or LOC, however, is still a long way off,
for a number of reasons. First of all, the ability to work with
real samples from start to finish in a microfluidic device,
in a hands-off sort of way, is usually very difficult. This is
because many “real” samples are inherently incompatible
with small channels, due to the presence of large particles or
wall-adsorbing molecular species that could lead to clogging.
The other issue holding back the development of sample pre931
treatment on chips is the sheer variety of samples with which
the chemist or biologist is confronted. There will be no one
universal approach to solving this problem on microfluidic
devices, just as there has been no universal solution in the
conventional chemistry laboratory.
Second, if one defines the LOC to be an entity that should
be able to perform all chemical functions and detection in
a monolithic device that fits in the palm of your hand
(or smaller), then existing devices are generally still quite
underdeveloped. This miniaturization will require increased
integration of not only fluidic elements, but also electrical,
optical, or other types of elements into microfluidic devices.
The MEMS world has amply demonstrated the integration of
mechanical and electrical functionality into small structures
for diverse applications. The aim of this review is to examine
more closely the role that MEMS technologies have played
to date not only in the fabrication of microchannels in
different substrate materials, but also in the integration of
electrical function into microfluidic devices.
II. PASSIVE MICROFLUIDIC ELEMENTS
The most basic elements of a microfluidic device are the
microchannels and microchambers themselves. These are
passive fluidic elements, formed in the planar surface of
the chip substrate, which serve only to physically confine
liquids to nL- or pL-volume cavities. Interconnection of
channels allows the realization of networks along which
liquids can be transported from one location to another on a
device surface. In this way, small volumes of solution may
be introduced from one channel into another, and controlled
interaction of reactants is made possible.
Analytical chemists turned to microfabrication techniques to be able to design and construct the microchannel
manifolds required to satisfy the very stringent volume
demands of TAS. This technology, the foundation on
which the MEMS field has been built, includes basic IC
techniques, namely, film formation, doping, lithography,
and etching, developed four decades ago for the microelectronics industry. In addition, special etching and bonding
processes that permit the sculpting of 3-D microstructures
with micrometer resolution have been and continue to
be developed [9]–[12]. Micromachining technologies are
primarily silicon-based, due both to the traditional role
of this semiconductor in IC technology and its excellent
mechanical properties [13]. Silicon is thus unique, as it
makes the combination of mechanical and electrical function
in single devices possible, providing the impetus for the
enormous activity over the past almost three decades in the
area of MEMS. In a parallel development, the high precision
obtainable with micromachining processes has led to their
application in the patterning of materials other than silicon.
Thus, ceramics, plastics, quartz, and glass are gradually
becoming accepted complementary materials to silicon,
widening the potential range of applications of microfabricated components and systems. This section considers
some examples of micromachining methods that have been
used for the formation of micrometer-dimensioned fluidic
channels in various substrate materials.
932
A. Microchannels in Silicon
Silicon micromachining can be divided into two categories, depending on what region of the wafer is being
worked. Bulk micromachining refers to those processes
which lead to structures in the single-crystal wafer, whereas
surface micromachining results in structures located on
the wafer surface. Bulk micromachining methods fall into
four categories, namely wet isotropic, wet anisotropic, dry
isotropic, and dry anisotropic [11], [12]. Isotropic methods
are those characterized by etch rates which are more or
less equal in all directions. Anisotropic methods, on the
other hand, have etch rates which are significantly faster
in a particular direction or directions. Wet etching involves
dipping the silicon wafer into a solution containing the
etchant. The etchant may etch certain crystal planes faster
than others (e.g., KOH, tetramethylammonium hydroxide).
In this case, the side walls of the resulting structure are
defined by the slowest etching crystal planes. For wafers
of 100 orientation, for example, channel geometries are
trapezoidal, with side walls corresponding to the 111
planes. For 110 wafers, the different orientation of the
111 planes makes deep, rectangular channels possible.
The isotropic wet etching solution most commonly used
is known as HNA, and contains hydrofluoric acid (HF),
HNO , and acetic acid. The resulting channels have rounded
side walls, and are wider than they are deep, since etch rates
are equal for all crystal planes. A discussion of the further
intricacies of silicon micromachining using wet etching
techniques is beyond the scope of this review. Ample
detailed examples are given in [9]–[12].
Reactive-ion etching (RIE) processes are dry, relying on
external RF power applied to a pair of plates to accelerate
stray electrons in the gas between them. The kinetic energy
of the electrons is augmented in this way to levels that allow
chemical bonds in the reactant gases to be broken, leading
to the formation of reactive ions and radicals. The wafer
to be etched is placed between the plates to be exposed to
this reactive plasma. Depending on the conditions, etching
can be achieved chemically, by reaction of fluorine free
radicals with the surface, for example, or physically, through
bombardment of the surface with high-energy ions [11],
[12]. These processes can be fully or partially isotropic
or anisotropic. One type of RIE, termed deep reactive-ion
etching (DRIE), has gained in popularity in recent years.
DRIE is a very high-aspect-ratio etching method for silicon,
with aspect ratios of up to 30:1 (height:width) possible.
This is because the plasma is highly dense. In addition, the
reactive ions are accelerated by applying a bias voltage to the
wafer, so that they bombard the wafer surface vertically. As
a result, very thin, tall structures and trenches can be formed
with this technique, at typical etch rates of 2–3 m/min
[14]–[16].
Microfluidic channels in silicon made by wet or dry
etching processes are generally sealed using a glass cover
chip, since this facilitates visual interrogation of fluids
within the device. Bonding in this case is accomplished by
field-assisted or anodic bonding [11], [12]. This involves
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
applying a voltage of between 200 and 1000 V over the
silicon/glass assembly, with the cathode (negative electrode)
on the glass side. The components to be bonded are heated at
the same time to a temperature between 180 C and 500 C.
Under these conditions, the glass becomes more conductive,
and Na ions in the glass migrate more rapidly toward the
cathode (that is, away from the glass–silicon interface).
This results in a large electric field at this interface, which
pulls the two surfaces together. Covalent bonds between the
silicon and glass are formed due to the elevated temperature
and applied electric field, although the mechanism is still
not so clearly understood. Pyrex 7740, a borosilicate glass,
has been the material of choice for anodic bonding. This
is because its thermal expansion coefficient nearly equals
that of silicon. The use of glues or other adhesive layers
to seal devices is not so common, as these approaches are
generally not so clean and can result in contamination or
even clogging of channels.
It is not surprising that some of the first microfluidic channels were made in silicon [17]–[19], given the role that silicon
plays in MEMS. (In fact, the 2-in silicon gas chromatograph
developed by Terry et al. [17] predated the introduction
of the TAS concept by 11 years.) In [18], standard etch
techniques were used to form channels in 100 -oriented
monocrystalline silicon, involving anisotropic etching using
either KOH or ethylenediamine/pyrocatechol. However, the
use of silicon is generally incompatible with electroosmotic pumping (EOP), which quickly became the method
of choice for liquid transport in microfluidic devices in the
1990s (see preceding text). Typical linear flowrates fall in
the range of micrometers per second to several millimeters
per second, with the required electric fields usually quite
high, corresponding to hundreds of volts per cm. Channel
lengths on the order of mm to cm thus necessitate applied
voltages which often exceed 1000 V. The substrate used
must therefore have excellent electrical insulating qualities.
Since silicon itself is a semiconductor, its usage is limited by the quality of the insulating layers of silicon oxide
and/or nitride that can be deposited on the surface. Generally
speaking, even good quality, pinhole-free insulating layers
with thicknesses up to 1 m suffer dielectric breakdown
at applied voltages of 720 V or less [18]. Silicon has thus
generally not been applied for electroosmotically controlled
microflow manifolds.
There are a few notable exceptions in the literature,
however, where silicon-based devices were used successfully in conjunction with low field strengths ( 100 V/cm)
for electrokinetic separations. These include demonstrations of free-flow electrophoresis of amino acids [20] and
proteins [21] [see Fig. 3(a) and 3(b)], and DNA analysis
by synchronized cyclic capillary electrophoresis [22] and
gel electrophoresis [23], [24]. Recently, Petersen et al.
reported microchip electrophoretic separation of caffeine,
paracetamol, ketoprofen, and ascorbic acid in a silicon
channel sealed with glass, using an electric field strength
of 550 V/cm over a channel 3.5 cm long [25]. In this case,
the silicon was coated with a waveguiding layer consisting
of 15 m of thermal oxide, followed by 6 m of silicon
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
oxynitride, and finally 4 m of undoped silica. Channels
(12 m deep) were etched into this layer by RIE, leaving a
13- m-thick thermal oxide layer between the channel and
the silicon substrate.
Pressure-driven flow applications have also been implemented in silicon chips, notably liquid chromatography (LC)
[see Fig. 3(c)–3(e)], [26]–[31], optical detection cells [32],
[33], microreactors [34]–[37], and miniaturized flow injection analysis ( FIA) [38], [39] (see also Section III-B3).
Examples of gas chromatography (GC) chips may also be
found in the literature [17], [40]–[43]. When high-aspect
1 channels in silicon are desired,
ratio depth:width
DRIE is the method of choice. Daridon et al. used DRIE
to fabricate a microflow system for ammonia analysis in
glass/silicon/glass devices [44]. Deep, narrow channels
30 m wide and 220 m deep made up the first part of the
silicon-based network, in which reagents were added to the
sample zone. This was done in a side-on fashion, yielding
two 15- m-wide curtains of solution running side-by-side
down the channel. Diffusion distances of sample and reagent
molecules were reduced in this way, allowing complete
mixing of solutions in less than 1 s by diffusion alone. To
be able to perform absorbance detection of the chemically
converted ammonia compound, a small-volume, long-pathlength optical cuvette was required. A small-diameter hole
was etched by DRIE through the silicon wafer to form this
element, which as a result had an optical pathlength equivalent to the wafer thickness. A recent publication by Juncker
et al. describes a microfluidic capillary system formed by
DRIE, designed to autonomously transport liquids from
one end to the other by capillary forces [45]. The principle
was demonstrated using a heterogeneous immunoassay
for a cardiac marker, which could be performed within 25
min. DRIE has also been used to fabricate surface-area
enhancing structures in silicon microchannels for extraction
and catalyzed reactions, in the form of arrays of staggered,
narrow silicon columns (see also Section III-B2). These
replace packed-bed designs, in which a channel is packed
with particulate material. In one example, the columns were
used to extract DNA from solution in a sample pretreatment
step [46]. Multiphase microreactor devices have also been
based on this principle [34]. Of course, arrays of small
holes through a wafer are also possible, as Thiébaud et al.
demonstrated in a device made for electrophysiological
measurements of slices of rat brain [47]. Diffusion of
nutrient medium from a reservoir under the chip to the cells
on top of the chip was assured in this way.
B. Microchannels in Glass and Quartz
Glass is a much better material, electrically speaking,
for microfluidic devices utilizing electrokinetic sample
transport. This, combined with the fact that it is optically
transparent, has made it one of the main materials used in
microfluidics today. Pyrex 7740, a borosilicate glass found
in many cleanrooms, has been popular for glass microfluidic
devices. From a fabrication point of view, however, there
has been no limitation noted so far as to the type of glass
933
Fig. 3. (a) Free-flow electrophoresis device made in silicon and sealed anodically with glass. The
photograph was taken through the glass cover. 1 marks the separation bed (50 mm long, 10 mm wide,
50 m deep); 2, the array of 125 inlet channels; 3, the electrode chambers (50 mm long, 2 mm wide,
50 m deep); 4, the arrays of 2500 channels separating electrode chambers from separation bed; 5,
the array of 125 outlet channels; 6, the sample inlet; 7, the buffer inlets; 8, the buffer outlet; 9,
common waste. (b) Scanning electron micrograph (SEM) of a corner of the separation bed where
both inlet channels (left) and 2500-channel array (right) enter it. The inlet channels are 70 m wide
(across the top) and 50 m deep; the channels to the electrode chamber are 12 m wide (across
the top), 10 m deep, and 1 mm long. (c) Assembled silicon/glass chip for small-volume liquid
chromatography. The separation channel (long horizontal channel) is 20 mm long, 300 m wide, and
100 m deep (490 nL); the injector is the channel element on the right; the detector cell (2.3 nL,
1 mm long, designed for absorbance detection) is at the end of the separation channel on the left.
(d) SEM showing the end of the separation column and entry into the detector cell. Two arrays
of 58 fine channels serve as a frit to retain particles in the separation channel. Note that the dead
volume between the end of the column and the cell is about 140 pL, so that this dead volume is
minimized. (e) SEM of a detail of the frit. These channels were 3.2 m wide, 1.4 m deep, 1.6 m
apart, and 19 m long (volume of each channel: 59 fL). It was possible to retain particles larger
than 3 m in the column with this barrier.
934
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 4. (a) Schematic diagram of the process flow used for glass micromachining in the authors’
laboratory. This process differs from others in that a polysilicon etch mask is used, rather than a metal
mask. (b) SEM of a double-T intersection etched in glass, used to perform volume-defined injection
in microchip electrophoresis (see Fig. 1). Channels are about 50 m wide across the top and 20 m
deep. The intersection is 200 m long, allowing the definition and injection of 165-pL sample plugs.
(c) SEM showing a channel cross section, again about 20 m deep and 50 m wide at its widest
point. The channel was thermally sealed with a second glass wafer. Note that there is no visible
bonding interface, indicating that the wafers have been hermetically sealed. (d) View of one full
FIA structure and partial views of two others. These devices consisted of three layers of glass,
with interconnecting vias and holes made by spark-assisted etching. Channels were 200 m wide
and 30 m deep. The networks are relatively complex, allowing reagents to be added to a sample
stream, and sufficient time for reaction before passage through an integrated optical cuvette. (Photo:
A. Daridon, IMT, University of Neuchâtel, Neuchâtel, Switzerland).
that can be used, as long as it is available in the form of
smooth, planar substrates. Microfluidic devices made in
fused silica (quartz) have also been reported. This material
is attractive because it is UV-transparent, a characteristic
which makes it superior to glass. Photolithographic processes similar to those used for silicon can be employed
for the patterning and subsequent transfer of channels into
planar glass and quartz wafer surfaces.
The glass micromachining method established in our laboratory is shown in Fig. 4(a). This is based on a process first
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
developed by Grétillat et al. [48] and has become standard in
our work [49]–[51]. Channels are etched into a 10-cm Pyrex
7740 wafer using HF, using a 200- or 400-nm-thick layer
of polysilicon as a masking layer over areas where etching
is not desired. A typical process flow would proceed as
follows. After deposition of the polysilicon by low-pressure
chemical vapor deposition (LPCVD), wafers are dehydrated
at 200 C for 30 min and exposed to hexamethyldisilazane
(HMDS) vapor for 15 min to improve photoresist adhesion. A 1.8- m-thin layer of AZ1518 (Clariant, Muttenz,
935
Switzerland) positive photoresist is then spin-coated onto
the wafer at 4000 r/min for 40 s and prebaked on a hotplate
for 1 min at 100 C. The resist layer is then patterned
photolithographically by illumination through an optical
mask. Laser-plotted transparency films with a dot size of
7 m (DIP SA Repro, Lausanne, Switzerland), mounted on
blank 5-in glass plates, serve as photolithography masks
for rapid prototyping applications. When higher resolution
or fine structuring are required, Cr masks with 1- to 2- m
resolution are used. In the case of transparency films, the
exposure dose is increased to 60 mJ/cm , a value which
is 10% above the recommended value for the resist, to
compensate for the reduced transparency of the laser-plotted
films. Wafers are then developed for 1 min in AZ351
developer (Clariant), which has been diluted 1:4 with
deionized, 18-M (DI-18M) water. This is followed by
a rinse in DI water and postbake at 125 C for 30 min.
RIE is used to transfer the pattern into the polysilicon
layer. After photoresist removal with acetone, the channel
structures are etched into the glass using 49% HF. Finally,
the polysilicon layer is removed by dipping the wafer for
5 min in 40% KOH solution at 60 C. The channels are then
thermally sealed with glass cover-plates containing holes
to access the microfluidic network. Once these holes have
been aligned with the ends of the corresponding channels
in the lower wafer, the two-wafer assembly is placed in
an oven for thermal bonding at 650 C. After an initial
temperature ramp, the wafers are held at 650 C for a couple
of hours. During this time, the wafers are fused together.
A slow cooling process is then initiated to prevent thermal
stress and the resulting cracking or breaking of wafers. A
thorough cleaning procedure to remove both organic and
inorganic residues (e.g., micrometer-sized particles from
the hole-drilling process) from the wafers is critical for
achieving good, void-free bonding. Fig. 4(b) shows a SEM
of a 200- m-long double-T junction made in glass, so
called because each of the side channels forms a T-shaped
intersection with the horizontal channel. An example of the
hermetic seal between two wafers that is typically achieved
with thermal bonding is given in Fig. 4(c). Note that there is
no visible interface between the wafer surfaces.
Glass etching with HF is isotropic in nature. The resulting
channels have rounded side walls, and widths which are
twice the channel depth plus the width of the initial line on
the etch mask [see Fig. 4(b) and 4(c)]. Channel depths are
determined by etch times. Typical dimensions are widths
and depths of tens of m, with channel lengths on the order
of a few cm. The etchants used in glass microfabrication
may vary in composition [52], [53], with mixtures of HF and
HNO [54], NH F-buffered HF [55], or concentrated [56]
or dilute HF baths being used. Metal etch masks composed
of a thin layer of Cr covered by Au are most common [54],
[56]. The Cr layer acts as an adhesion layer to better coat the
Au onto the glass surface. Polysilicon [48] and amorphous
silicon [56] may also be used as etch mask layers, as described previously. For very shallow structures, photoresist
alone may be adequate to protect the glass surface, although
it quickly deteriorates in the more aggressive etch solutions
936
[55], [56]. Quartz devices have been fabricated using similar
techniques as those described previously, though the thermal
bonding must be carried out at higher temperatures, around
1000 C or so [57]–[59].
Alternative glass-to-glass bonding methods are available.
For instance, anodic bonding of glass to glass via an intermediate, 160-nm layer of silicon nitride at a temperature
of 400 C and a voltage of 100 V has been reported [60].
Low-temperature glass-to-glass bonding is possible, using a
1% HF solution, wicked in between the wafers, and pressure
(1 MPa) at room temperature [61]. Another low-temperature
approach involves a spin-on layer of sodium silicate solution
as adhesive layer [62], [63]. Spontaneous bonding of thoroughly cleaned and hydroxylated wafers at room temperature has also been described for both quartz and glass [64].
Generally speaking, channels are formed in one layer and
sealed by a second. It is also possible to achieve symmetrical
channel cross sections by mating two surfaces containing
channels [51], [65]. The inclusion of special alignment marks
on the wafers to be bonded can yield an alignment precision
of 5 m, dictated only by the optical resolution of the
mask aligner used [65]. Three-layer glass devices for FIA,
incorporating through-holes from one layer to the next, and a
200- m-diameter optical cuvette formed through the middle
wafer, have also been reported [51]. These were made and
bonded in the same way as shown in Fig. 4(a). The challenge
in this case was the formation of holes small enough not to
cause dispersion of the sample zone as it passed from one
level to the next. Particularly critical in this regard was the
fabrication of the optical cuvette. Spark-assisted etching,
also known as electrical discharge machining, proved to be
a valuable tool for making holes as small as 200 m in
diameter through the 525- m glass wafers. Fig. 4(d) shows
several bonded devices still in wafer format, before dicing
(cutting) of the individual chips. Multilayer glass devices
with 3-D microchannel networks for chemical synthesis
applications have also been reported by Kikutani et al.
[66], [67]. The “pile-up” glass microreactor described in
[67] contained ten layers of microchannel circuits, which
were thermally bonded all at once in much the same way
as described previously.
RIE of quartz has been employed to produce a liquid
chromatographic chip with an array of 10- m-high,
5- m 5- m rectangular columns to act as support for
the stationary phase in the 4.5-cm-long, 150- m-wide
separation channel [68]–[70]. Fig. 5 is a SEM of this type
of device, taken at the entrance to the separation column.
It clearly shows how the inlet channel is bifurcated several
times to ensure even solution distribution over the array
of diamond-shaped pillars at the top of the photo. Chromatographic performance was good [68], and peptides
stemming from the tryptic digest of ovalbumin could be
separated [70]. DRIE of Pyrex has recently been well
characterized, with a 20- m-thick nickel mask being used
to achieve good verticality and aspect ratios greater than
10 [71]. Ceriotti et al. also used a nickel mask to fabricate
50- m-deep rectangular channels in quartz by inductively
coupled plasma (ICP)-RIE, using the process flow shown
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 5. SEM of structure designed for microchip chromatography,
showing channel layout at the beginning of the column. All features
have been etched by RIE to a depth of 10 m. (Reprinted in part
with permission from [68]. Copyright 1998 American Chemical
Society).
in Fig. 6(a) [72]. Detection of fluorescently labeled amino
acids separated electrophoretically in a DRIE quartz channel
much like that in Fig. 6(b), sealed with an elastomer chip,
was demonstrated [73].
Photostructurable glasses such as FOTURAN (Schott)
are also interesting materials for TAS, as they allow for
microstructures having aspect ratios of 20:1, and the possibility of transformation to a glass ceramic upon heating [74].
These materials are inherently photosensitive, and hence do
not require a photoresist layer for patterning. Exposure to
UV results in the formation of microcrystallites, which can
subsequently be removed by etching in HF. Tips for atomic
force microscopy have been fabricated in this material [75].
Channels having a minimum depth of 25 m are also possible [74]. Recently, Becker et al. reported the fabrication
of a microchip electrophoresis device in a new type of
photostructurable glass [76]. A separation of fluorescently
labeled amino acids was demonstrated in channels which
were 500 m deep and 200 m wide.
C. Miniaturized Transparent Insulating Channels ( TICs)
There are a number of alternative approaches for the fabrication of insulated microchannels which still use silicon.
One of these involves the formation of channels with extremely thin walls ( 1 m) in silicon nitride or oxide, using
a thick ground plate to support these structures [77]–[80].
The process used to produce the channels is shown in Fig.
7(a). To start, channels are patterned and etched in silicon,
using either wet or dry, anisotropic or isotropic methods.
The structure on the right in Fig. 7(a) has a geometry which
is characteristic for an isotropic process. The middle and left
channels, on the other hand, were formed with anistropic
processes. (Note that the combination of all these channel
geometries in one Si wafer is unrealistic. The geometries
are given to indicate that there is some flexibility in the
choice of profile geometry available when using silicon.
This is in contrast to glass, for which anisotropic processes
are not yet well established.)
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
Fig. 6. (a) Process flow for ICP-RIE of 50-m-deep rectangular
channels in quartz. (b) SEM showing the cross-sectional profile of a
dry-etched channel in quartz. Width across the top: 50 m. Depth:
about 50 m. Conditions for etching had not been optimized in this
case. It was possible in later experiments to achieve good wall
verticality and smooth walls [72].
Once formed, the masking material used to form the
channels is removed from the silicon surface, and the channels are coated with an insulating layer. The silicon wafer is
then anodically bonded via the insulating layer to a Pyrex
wafer. After bonding, the silicon is etched away from the
937
backside using an isotropic etchant, leaving the freestanding,
thin-walled microchannels. An early publication described
the deposition first of a 50-nm-thick layer of silicon nitride,
followed by an oxide layer of up to 600 nm [78], to facilitate
bonding to the Pyrex. Later work used only silicon nitride
layers of 360 nm [79] or 390 nm [80] to form the channels. In
any case, these channels are electrically insulating, optically
transparent, and demonstrate very efficient heat dissipation,
making them ideal candidates for electrokinetically driven
microfluidic applications. Fig. 7(b) and (c) shows SEMs of
TICs molded in an isotropically etched Si channel, and
a DRIE channel, respectively. As is evident in Fig. 7(c),
TICs do have the disadvantage that they are fragile. This
may be overcome by covering the channels with a glue or
other polymer layer [77]–[79]. This type of structure has
been used for microchip electrophoresis of small cations in
conjunction with conductivity detection [79], and for the
demonstration of field-effect flow control in microfabricated
structures [80].
D. Microchannels by Replication in Poly(Dimethylsiloxane)
(PDMS)
Poly(dimethylsiloxane) (PDMS) is a polymer which is becoming increasingly popular for microfluidic applications,
because structures made in this material are inexpensive,
easy to handle, and rapidly fabricated by replica molding
under noncleanroom conditions. The special properties of
this elastomer, and its use in soft lithography (replication)
and micro contact printing (surface patterning by deposition
of molecules using stamping techniques) to obtain micrometer-sized structures, are described in several reviews
[81]–[83]. The use of microfluidic devices replicated in
silicone rubber was reported as early as 1989 by Masuda
et al. for a biological application involving cell fusion
[84]. Fig. 8(a) is a schematic diagram of a typical casting
procedure for replication of microchannels in PDMS.
Fabrication of microchannels involves casting a solution
of the PDMS prepolymer onto a master whose surface
has been structured to yield a topography or surface relief
of some kind. When cured, PDMS faithfully replicates,
with nm resolution, the surfaces with which it has been in
contact. Microchannels are thus easily formed in PDMS if
the corresponding master has a raised network of ridges to
serve as microfluidic network mold. PDMS has a number of
intrinsic properties which make it interesting for microfluidic applications. It is, for example, optically transparent in
the UV and visible, from 230 to 700 nm [72], [85]. The gas
permeability of PDMS means that bubbles created inside
channels by electrolysis of water or from other sources may
be dissipated through the material [86]. Gases can of course
also penetrate the PDMS to enter microchannel structures,
making it a suitable material for applications using live cells
[83], [85]. Structures with flat surfaces made in this material
seal reversibly with glass, silicon, and PDMS. Alternatively,
irreversible bonding can be accomplished by treatment of
the surfaces to be bonded with an oxygen plasma [87], [88].
A variety of masters have been used for replication of
microchannels in PDMS. Anisotropic wet etching of 100
938
Fig. 7. (a) Basic process used for the fabrication of TICs.
Cross-sectional views of TICs made using (b) an isotropically
wet-etched silicon mold and (c) a DRIE silicon mold. [(b) Reprinted
in part with permission from [78]. Copyright 1998 Elsevier Science
Ltd.; (c) Reprinted in part with permission from [79]. Copyright
2001 Wiley-VCH Verlag GmbH].
silicon has been used to produce masters for plastic channel
replication [89]–[91], but the resulting channels had trapePROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 8. (a) Schematic diagram of the steps involved in replication of microchannels in PDMS. (b) A
three-layer PDMS device for passive mixing by lamination of solution streams. (Reprinted with
permission from [85]. Copyright 2000 IEEE) (c) A 3-D PDMS network for a cell culture application.
(Reprinted with permission from [100]. Copyright 2002 IEEE.)
zoidal cross sections with aspect ratios 1. There is little
freedom in channel network design, since possible layout
geometries are dictated by crystalline orientation. To retain
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
the freedom of channel layout afforded by isotropic etching
of glass, a nickel master can be prepared directly from a
structured glass wafer by growing a relatively thick (3-mm)
939
nickel layer onto it [92], [93]. Alternatively, resist reliefs on
silicon wafers made using the negative photoresist, EPON
SU-8, can be used as masters. This approach is an attractive choice in terms of fabrication time and layout flexibility
[85], [87], [94], [95]. Moreover, features 5 to 1200 m high
are possible with aspect (height-to-width) ratios approaching
20 [96]–[98]. To date, 1 : 1 aspect-ratio structures [negative
channel structures (i.e., ridges) with equal height and width]
were used for PDMS molding, and they can be used indefinitely [83]. Masters realized by DRIE have the same advantage as those made in photoresist, namely, freedom to choose
channel layout [88], [99]. Curved relief patterns with vertical
walls and high aspect ratios are thus possible.
3-D PDMS structures have been reported by several
groups. In one case, many thin ( 100 m thick) patterned
PDMS layers were stacked to form 3-D channel paths [85].
Each layer contained channels and openings, molded against
an SU-8 master using a sandwich-type molding configuration. Fig. 8(b) shows a photograph of an assembled 3-D
passive micromixer made in this way. Anderson et al. used
a similar approach yielding thin layers with through-holes
and microstructuring on both sides, which were sealed with
thicker PDMS slabs [95]. A microfluidic device for the creation of a sheath flow of buffer to hydrodynamically confine
a sample stream close to a sensor surface was realized in
PDMS by Hofmann et al. [99]. This approach required the
ability to introduce a buffer flow onto the sample stream
from above to compress it, a criterion which this 3-D device
fulfilled well. Fig. 8(c) shows a close-up SEM image of a
3-D microchannel network formed in PDMS for culturing
of human cancerous liver cells [100]. The 3-D approach
taken here could prove useful for more sophisticated tissue
engineering applications.
E. Microchannels by Replication in Other Polymers
Masters made in silicon can be used to imprint or hotemboss channels in hard plastic materials like poly(methyl
methacrylate) (PMMA) at temperatures close to the softening point of the plastic or, alternatively, at elevated pressures [101], [102]. Martynova et al. used the former approach
to make plastic microchannels [103], whereas Xu et al. were
able to do much the same but at room temperature and high
applied pressures [104]. McCormick et al. used a micromachined silicon wafer to fabricate a metal stamp by electroplating Ni onto the silicon, and using this electroform to
create a second electroform [105]. The second form is thus
a replica of the original silicon master. These researchers
used forms made using this two-step electroplating process to
both imprint and injection-mold channels into plastic. Other
reports of injection molding for microfluidic chips may be
found in [106] and [107]. Excellent discussions generally
about the fabrication of polymer microfluidic devices are
given by Becker and Gärtner in [101], and again by Becker
and Locascio in [102].
A more recent report describes the 3-D replication of structures in UV-curable acrylates using a finely patterned silicon
master, produced by a series of dry etching processes [108].
940
Though this technique has not been applied to microfluidic
devices yet, it offers a route to nanostructuring, and hence
could be an attractive tool for the microfluidics community.
F. Other Techniques for Fabrication of Polymer
Microfluidic Devices
Laser ablation was first introduced by Roberts et al.
for the fabrication of polymer microchannels [109]. This
technique involves directing laser pulses at the plastic surface in defined regions, which causes degradation of the
plastic at those spots as a consequence of a combination
of photochemical and photothermal degradation processes.
A UV excimer laser was used to produce channels in a
variety of substrates, which were then subsequently sealed
using a low-cost plastic lamination technique. EOF in these
channels was characterized, and shown to be a function of
ablation conditions [109]. Henry et al. studied this aspect of
laser-ablated channels in more depth, using channels made
in poly(ethylene terephthalate glycol) [110]. Johnson et al.
used laser ablation techniques to modify the surfaces of
channels already hot-embossed into various polymer substrates [111], [112]. In one case, the surface structures
were designed to reduce broadening of sample plugs as
they passed around corners [111], whereas in the other, the
purpose of the modification was to create a rapid mixer
[112]. CO -laser machining has also been used to rapidly
write structures in PMMA [113], and has proven to be an
effective tool for rapid protoyping of devices for diagnostic
applications [114].
A simple method based on plasma etching was recently
presented for producing both channels and electrodes in
polymer, starting with three-layer foil composed of 50 m of
polyimide sandwiched between two 5- m layers of copper
[115]. Photoresist is used to pattern the copper layers, after
which the exposed copper is etched away chemically using
an aqueous solution of CuCl –H O . The foil is then placed
in a reactive plasma chamber to transfer the desired pattern
into the polyimide. After the plasma etch, the copper is
either removed or further patterned to produce conductive
pads, which can be coated with Au by electroplating. The
process is shown in Fig. 9(a), along with some photos of
the finished devices in Fig. 9(b). Implementation of such
devices for microchemical analysis was demonstrated with
voltammetric detection in a 60-nL microchannel.
III. INTEGRATION OF ELECTRICAL AND ELECTROCHEMICAL
FUNCTION INTO MICROFLUIDIC DEVICES
One of the unique possibilities offered by MEMS technology is the direct integration into microfluidic devices of
elements not having a fluidic function per se. The fact that
these technologies were developed for the microelectronics
industry in the first place makes their use for fabrication of
features with electrical function rather obvious. The ability
to deposit and structure thin metal or other conductive films
on wafer or channel surfaces is thus being exploited in microfluidics for integrating electrochemical detection (ECD),
resistive heating, and pumping elements into devices. This
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 9. (a) Process flow for plasma etching microchannels in
polyimide foil. A) Channels are formed first, followed by a second
patterning/etching step to form access holes. It is during this step
that channels may be etched down to contact the underlying
Cu layer. B) This copper layer may be further structured to
yield electrodes and electrical contact pads. (b) Photographs
of a four-electrode device made in polyimide foil. (Reprinted
with permission from [115]. Copyright 2002 Royal Society of
Chemistry.)
section will consider examples which have an electrical element somewhere on or in the chip for performing one or
more of these functions.
A. Electrochemical Detection in Microfluidic Chips
ECD, which includes amperometric, potentiometric, and
conductometric detection, is the subject of an increasing
number of research papers looking at its integration into
microfluidic chips [116], [117]. This is because it scales
better upon miniaturization than absorbance or fluorescence
detection, since output signal is dependent on electrode
surface area rather than on available detection volume [118].
As a result, limits of detection (in concentration terms)
do not degrade as rapidly for electrochemical detection as
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
they would for optical techniques [119]. Electrochemical
techniques are generally attractive for this reason, though
they will never compete with fluorescence in terms of sensitivity. The use of micromachining technologies to integrate
electrochemical sensing elements into microfluidic devices
brings some unique advantages not possible in conventional
systems. For instance, fabrication of electrodes directly in
microfluidic channels means that these are aligned a priori,
eliminating the need for often-difficult alignment procedures. Moreover, devices incorporating detection electrodes
do not require a large amount of peripheral equipment for
acquisition of detector signal. This makes the development
of portable instrumentation based on microfluidic devices
with ECD a realistic possibility.
There have been a number of approaches reported in the
literature for integration of electrodes into chips. In the simplest case, grooves or channels whose purpose it is to simply
hold metal or carbon electrodes in place are formed in one
of the surfaces of the device, and sealed with the second
layer. Electrodes are inserted into these channels before assembly of the devices. This approach has been reported for
chips replicated in PDMS and utilizing 30- m carbon fiber
electrodes [120]–[122]. Pt wires 127 m in diameter were
inserted in 130- m guide channels for contact conductivity
detection in chips which had been hot-embossed in PMMA
[123], [124]. In [125], the end of an electrophoresis separation channel in glass was widened by etching in HF, to facilitate insertion of Pt, Au, or Cu wires for amperometric
detection. Alternatively, electrode channels were formed in
PDMS and filled with carbon paste [126], or laser-ablated in
polyethylene terephthalate (PET) and filled with carbon ink
[127]–[130].
More sophisticated means of integrating electrodes for
ECD involve deposition and patterning of metal layers
according to established micromachining techniques. The
technique used will depend on the type of metal chosen.
In the simplest case, glass devices for microchip electrophoresis were modified by cutting off the end of the chip
perpendicularly through the separation channel to sever the
waste reservoir from the chip. In this way, the outlet of the
separation channel was directly at the flat end of the chip,
rather than in a solution reservoir. Gold electrodes were then
formed around this channel outlet [131], [132]. In one case, a
thin gold film was sputtered around the outlet to a thickness
of 200 nm. Electrical contact was made using a lead glued
with silver epoxy to the gold layer. The gold electrode plus
lead was then coated with an electrically insulating ink,
leaving a gold disk-shaped area of 78 mm exposed for
use in amperometric detection [131]. In the other example,
electroless deposition of gold was carried out on the end of
the chip, using formaldehyde as a chemical reducing agent
to plate gold from solution onto the glass surface [132].
Again, the actual active electrode surface was defined when
the wire contact and gold surface were covered with an
insulating layer to leave only a small region bare.
In the previous examples, patterning using a photolithographic process was not possible, since the electrodes were
located on the edge of the devices, rather than on a planar
941
surface. More precise structuring of electrodes results when
photolithographic patterning is used. The simplest procedure
involves deposition of a uniform metal layer onto a planar
surface, followed by standard photolithography and transfer
of the electrode geometry in the metal layer. Generally, a thin
layer of one metal is first deposited to enhance the adhesion
of a second, thicker metal layer to be used as electrode. Typical adhesion layers for different metals include: 1) Cr (for
Au); 2) Ta or Ti (for Pt); and 3) Ti (for Cu). This was the approach followed by Martin et al. [133], who coated soda-lime
glass squares sequentially with chromium (5 nm) and gold
( 200 nm) using an evaporation technique. Chromium was
used as an adhesion layer for the gold. The Cr–Au layer was
then patterned using a positive photoresist, and the underlying Cr–Au layers removed in appropriate etchant solutions
in areas where the resist had been stripped during development. The result was an array of four parallel electrodes,
40 m wide and with a pitch of 80 m. Because these electrodes protrude slightly from the surface, the wafer surface
is no longer completely flat. Hence, hermetically sealing this
surface by thermal or anodic bonding to a second chip containing channels would not be possible if the second substrate
were glass or silicon. Martin et al. used a PDMS substrate instead, since this material conforms well to a surface bearing a
topography [133]. The PDMS and glass chips were aligned
in such a way that the electrodes were positioned perpendicularly to the separation channel, just beyond the end of
this channel. A similar procedure was used by Liu et al. to
produce a single-gold-electrode PDMS/glass device [134].
In a microreactor application, interdigitated arrays of carbon
microelectrodes were integrated into a silicon-based device
containing two flow chambers. The upstream chamber was
packed with enzyme-modified porous glass beads for glucose oxidation. The second chamber contained the electrode
arrays for electrochemiluminescence (ECL) detection of the
hydrogen peroxide produced, each consisting of 125 pairs
of 1-mm-long, 3.2- m-wide electrodes, with an electrode
spacing of 0.8 m. To obtain these detectors, the electrodes
were defined on a sputtered thin film of carbon with a standard photolithographic procedure, followed by structuring
of the carbon layer using an O plasma [135], [136]. The
flow chambers were defined after electrode fabrication by
photolithography in a thick layer (300 m) of Epon-SU 8
[135]. Woolley et al. presented a glass CE device with integrated Pt electrodes for indirect ECD of DNA [137]. As the
SEM image of this device in Fig. 10 shows, photolithography
makes possible the positioning of the 10- m-wide, dual-lead,
platinum working electrode just 30 m beyond the end of
the separation channel in the large reservoir. This is important, as this detection electrode finds itself just outside the
high electric field used for separation, and is effectively decoupled from this field as a result. In this case, 20 nm of Ti
(adhesion layer) and 260 nm of Pt were sequentially sputtered onto the substrate. Thick photoresist (ca. 10 m) was
then coated onto the wafer and patterned, and the exposed Pt
removed in hot aqua regia (personal communication with R.
A. Mathies, University of California, Berkeley).
942
An alternative way of structuring electrodes in microfluidic devices uses a microfabrication procedure known as
liftoff. This approach is more involved than that described
previously, requiring several steps. However, it remains
popular, particularly when making electrodes in platinum,
a metal which does not etch easily (see the previous text).
The liftoff process is described in more detail in Hierlemann
et al. (in this issue). Liftoff processes have been used to
form microelectrode arrays in microfluidic devices in gold
for generation of ECL [138], [139], and platinum, for
amperometric [135], [138] and conductivity [140] detection.
Three images of a microdevice incorporating two arrays
of Pt microelectrodes for glucose detection are shown in
Fig. 11 [138]. A microchip electrophoresis device in glass
incorporating a single platinum electrode for wireless ECL
detection has also been reported [141].
As mentioned previously, electrodes formed on top of
planar surfaces make good sealing with other planar substrates in hard materials difficult. A number of researchers
have developed a variation of liftoff processing which
circumvents this problem. The procedure starts much the
same, with a layer of photoresist being spin-coated onto a
wafer surface and electrode geometries defined. However, at
this point, shallow cavities ( 500 nm) are formed first in the
regions where electrodes are to be deposited. Only then are
the metal layers deposited onto the wafer, with thicknesses
that just fill the cavities in the substrate. The resist is then
removed as before by dissolution in an organic solvent,
with liftoff of the metal occurring at the same time. In this
way, the chip surface containing the electrodes retains its
planarity, and may be bonded to another silicon or glass
substrate. This approach was taken to form 160-nm-thick Pt
electrodes in a silicon chip for use in coulometric nanotitration in stopped-flow [142] and continuous-flow systems
[143]. Pt electrodes were made similarly in 200-nm recesses
in glass for conductivity detection [79], and 300-nm recesses
in glass for amperometric detection in conjunction with
microchip electrophoresis [144]. The device in Fig. 12 was
somewhat different, in that 170-nm-thick Pt–Ta electrodes
were formed in 12- m-deep cavities [145]. The application
in this case was contactless conductivity detection for
microchip CE. Good capacitive coupling into the detection
volume over a thin glass wall was thus required, while at
the same time ensuring complete isolation of the electrodes
from the solution in the analysis channel. Fig. 12 shows a
photograph of the finished device, focusing on the detector
region.
B. Integrated Temperature Control
There are a number of ways to generate heat in MEMS
devices, which have been developed for a variety of applications (see, e.g., Kovacs [11]). In microfluidics, the most
commonly implemented approach involves integration of a
simple resistor somewhere on the device. Passage of current
through a resistive element results in the generation of Joule
, where is electrical resisheat, , according to
tance and is current. Microfluidic devices are attractive for
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 10. Microchip electrophoresis device with integrated Pt counter and working electrodes
for amperometric detection. (A) Overall layout of the device. (B) Detailed schematic diagram of
the arrangement of electrodes at the end of the separation channel. A small Ag/AgCl reference
electrode was immersed in the detection reservoir to complete the three-electrode setup. (C) Close-up
SEM image of the photolithographically patterned Pt working electrode positioned just after the
end of the separation channel. (Reprinted with permission from [137]. Copyright 1998 American
Chemical Society.)
performing chemical reactions or processes requiring temperature control, since the high heat transfer rates in these
systems lead to easier implementation of uniform temperature conditions [146]. This section will examine microfluidic
devices incorporating resistive heating in one form or another
for both chemical and biochemical applications.
1) Biochemical Applications: There have been many
publications describing the application of MEMS technologies to the development of devices for DNA analysis.
A significant number have focused on the integration of
the polymerase chain reaction (PCR), used to increase the
amount of DNA from a sample for analysis, into micromachined chambers. Microchip PCR provides an attractive
alternative because the low thermal mass of these devices
allows the reduction of cycle times from several minutes to
30 s or less. Another DNA-related topic that has received a
lot of attention in recent years is the use of microchip gel
electrophoresis for DNA analysis (see, e.g., [147]–[149]).
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
Several of these use integrated heaters to achieve denaturation of double-stranded DNA [150], [151]. An in-depth
review of the general topic of nucleic acid analysis with
micromachined devices is presented by Mastrangelo (in this
issue).
Yamamoto et al. contacted a PDMS chip containing a microreactor array with a glass temperature control chip for
cell-free protein synthesis [152]. The glass chip had heaters
and sensors structured in a 500-nm-thick layer of indium tin
oxide (ITO) by etching in a 1:1 HNO :HCl solution at 50 C
for about 20 min using a patterned resist layer as etch mask.
Contact electrodes were then formed by liftoff, and the entire surface passivated by a 500-nm layer of SiO . Because
the ITO layer is optically transparent, optical detection was
possible through both chip surfaces. Thermal time constants
for heating and cooling were 170 ms and 3 s, respectively,
and uniform temperature gradients could be achieved across
each reactor. The cell-free synthesis of green and blue fluo943
Fig. 11. Three images of a microfluidic device designed to
function as an enzymatic microreactor. The devices incorporate two
arrays of Pt microelectrodes formed by liftoff. (a) Optical image
of empty devices before wafer dicing. The microreactor geometry
was defined in a 300-m-thick layer of EPON SU-8, and contains
two chambers separated by 11 posts. The chamber with the zigzag
shape was packed with glass beads that had glucose oxidase
immobilized on them. These were retained in this chamber behind
the posts. The H O produced when a glucose-containing sample
was reacted with the beads could pass the posts, to be detected
amperometrically at the electrode arrays, visible as light-reflecting
squares in this image. (b) SEM of a single microreactor after dicing.
The two electrodes arrays are more visible in this image (upper left).
(c) Close-up view of the posts and an electrode array, obtained by
scanning electron microscopy. The Pt electrodes are 3.2 m wide,
0.8 m apart, and about 1 mm long. [Fig. 11(a) and (b) reprinted
with permission from [138]. Copyright 2000 Kluwer Academic
Publishers; photographs courtesy of G.-C. Fiaccabrino, IMT.]
rescent proteins from their DNA templates was successfully
demonstrated.
Controlled protein crystal growth was the objective
of studies carried out by Sanjoh et al. with the help of
microfluidic silicon devices [153], [154]. The microfluidic
944
devices typically contained a crystal growth cell, reservoir
cell, and, for certain studies, heaters. The lower surface of
the growth cell was patterned with fine features in either nor p-type silicon layers, which serve as spatially selective
nucleation sites by electrostatically attracting dissolved
protein molecules. The integrated heaters were used to
control the degree of supersaturation of protein in solution.
Lysozyme crystals grown on n-type Si features are shown
in Fig. 13. In Fig. 13(a), the features are narrow ridges, as
shown in the inset, whereas in Fig. 13(b), they are trenches.
Crystallization is well-defined and clearly selective for the
n-type Si.
2) Microreactors for Chemical Engineering: Miniaturization, and hence MEMS technology, is attractive to the
chemical engineering community for the realization of microfluidic platforms for chemical process optimization. Of
particular interest is the capability MEMS offers to integrate
control, sensing, and reactor functions into the same device.
It has also been proposed that reactions, once optimized,
could be scaled out in arrays of microreactors rather than
scaled up to larger volumes in a single reactor. This would
certainly reduce the risks associated with handling particularly explosive or exothermic reactions, making chemical
production safer. It also becomes conceivable to think of
on-site generation of hazardous chemicals, thereby forgoing
the need for special storage and transport conditions [146],
[155].
Improved temperature control is ultimately one of the
main advantages to be gained from downscaling. Microfabricated microreactors for chemical engineering purposes thus
have often included elements for heating and temperature
sensing. A very simple example by McCreedy et al. saw the
insertion of a Ni–Cr wire, twisted into a serpentine shape,
into a PDMS solution before it was cured, to serve as a heater
in the resulting polymer chip [156]. This chip was sealed to
a glass base-plate that contained the microreactor channels.
Sulfated zirconia was used as catalyst, and was immobilized
in the inner surface of the PDMS cover. The PDMS was
stable up to 180 C, and it was possible to dehydrate hexanol
and ethanol to hexene and ethene, respectively.
Srinivasan et al. described a more sophisticated, Si-based
microreactor [37]. Pt–Ti heaters and sensors for temperature and flow were formed by liftoff on a 1- m silicon
nitride layer deposited on silicon. The T-shaped reactor
chamber was then etched from the other side down to the
nitride membrane, and a thin Pt catalyst film deposited by
shadow-mask evaporation on the inside surface of the membrane. The highly exothermic, Pt-catalyzed partial oxidation
of ammonia was considered, and the safe investigation of
ignition-extinction behavior demonstrated. Pt–Ti heaters
and resistance temperature detectors (RTDs) were incorporated by the same group into a Pd membrane reactor, for
purification of hydrogen, or alternatively, for hydrogenation
and dehydrogenation reactions [146]. The gas–liquid–solid
reaction of cyclohexene with H in the presence of a Pt
catalyst was chosen as the model reaction in another set
of three-layer microfluidic devices [34]. Two structured
layers of silicon were topped off by a glass cover-plate, onto
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
Fig. 12. Layout of a microchip electrophoresis device designed for small-ion analysis using
contactless conductivity detection. The inset on the right shows a photograph of the detector region.
Electrical connections to the electrodes were made via round contact pads at the end of the metal
lines. All metal structures were placed in 12-m-deep recesses to prevent problems during thermal
bonding due to uneven surface topography. (Reprinted with permission from [145]. Copyright
2002 Wiley-VCH Verlag GmbH.)
which a Pt–Ti thin-film heater and RTD had been made
[see Fig. 14(a) and (b)]. These contained either a packed
bed of catalytic particles or structured microchannels with
staggered arrays of columns, 50 m in diameter, 300 m
tall, with nearly 20 000 columns in all, to act as catalyst
support [see Fig. 14(c)]. It was pointed out that while the
high thermal conductivity of silicon allowed excellent heat
dissipation and hence good thermal control by the heaters,
large amounts of heat were lost to the environment. In
addition, heater performance was very dependent on how
the chip was packaged.
3) Other
Microchemical
Reaction
Applications: Classical wet chemical analytical methods are
generally based on the chemical conversion of the analyte
of interest through derivatization, complexation, or other
means to make it more visible to a detector. Many of
these methods lend themselves well to automation by
flow injection analysis (FIA). In this technique, sample is
injected into a flowing carrier stream and subjected to
various sample handling steps, such as dilution, reagent
addition, mixing, and finally, detection. Reaction times are
defined by the residence time of the reacting sample zone in
the flow system, and hence can be adjusted to some extent
by flow-rate variation. Precise flow control is crucial to
obtaining reproducibility in FIA, since measurements are
done under nonequilibrium conditions.
A number of publications have described the integration
of classical wet chemical methods into micromachined fluVERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
idic systems. One early example using a stacked system of
silicon chips 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 [38],
[157]. This example was interesting in that it was the first to
use silicon-based micro pumps for controlling liquids within
the valveless system. However, a smaller flow system did not
mean enhanced reaction, since the kinetics of the reaction
used for phosphate analysis are slow and proved to be the
limiting factor for analysis. A more recent example of FIA,
using much smaller chips, was developed for analysis of ammonia in waste and drinking waters. It used the Berthelot reaction to convert ammonia through reaction with phenol to
a blue indophenol dye. Again, the kinetics of this reaction
are slow, with maximum absorbance achieved only after 15
minutes or so at 25 C. Since a sensitive measurement was
desired in 5 min or less, and because reaction rates increased
significantly at 35 C and 45 C, the three-layer microfluidic device used had a 50-nm-thick ITO layer sputtered on
one surface for resistive heating [44]. An all-glass version of
the three-layer chips for this application, complete with ITO
layer, has also been reported [51].
Fast derivatization is especially important when analytes
are to be labeled on-line just before or after a separation.
Eijkel et al. micromachined a heated chemical reactor for
precolumn chemical derivatization of amino acids with the
fluorescent agent, 4-fluoro-7-nitrobenzofurazan (NBD-F)
before separation in a conventional high-performance
945
Fig. 13. Photographs show lysozyme crystals that have been
grown in a growth cell with n-Si features patterned in the bottom.
(a) The n-Si nucleation sites take the form of ridges, whereas in (b),
they are narrow trenches. In both cases, crystals clearly prefer to
form on these sites. Solution pH: 4.7. (Reprinted with permission
from [153]. Copyright 1999 Elsevier Science Ltd.)
liquid chromatography system (HPLC) [158]. The device
contained on-chip Pt–Cr thin-film metal resistors as heaters,
which were recessed into the silicon chip by sputtering the
metals into a 1.5- m-deep groove. These elements could
heat a 50- L volume to a maximum of about 90 C at a
rate of 2 C/s, using 2 W of power. In addition, there were
two temperature-sensitive resistors on board for temperature
measurement.
4) Other Examples Using Integrated Temperature Control: Analysis of gaseous analytes often requires their preconcentration by adsorption onto a solid surface, followed by
a thermal desorption step once enough sample has been collected for further detection. Microfluidic devices designed
for this purpose and incorporating thin-film resistive heaters
have been reported. In one case, airborne benzene, toluene,
ethylbenzene, and xylenes were of interest [159]. A Pt heater
was formed by sputter deposition and liftoff on the bottom
of a Pyrex wafer, which then had a trench machined into it
and was sealed by anodic bonding to a second Pyrex wafer.
The trench was packed with adsorbent particles, and gaseous
samples passed through the packed bed. Gases preconcentrated in this device were desorbed by ramping the temperature at about 2 C/s, and collected in a second micromachined
detection cell. A preconcentration factor of 100 could be obtained this way, and parts-per-million (ppm) levels of toluene
could be determined. The same researchers (Ueno et al.) later
946
Fig. 14. (a) Schematic overview of a three-layer silicon–glass
device for multiphase mixing and reaction. The thin-film metal
heater and temperature sensor pattern is superimposed onto the
channel layout. (b) Photograph of the device, corresponding to the
schematic in (a). (c) SEM of a small portion of the 20 000-column
array in the reactor bed, each of which is 300 m high and 50 m
in diameter. These structures serve as catalyst supports within the
microreactor. (Reprinted with permission from [34]. Copyright
2002 IEEE.)
used this device for measurements of mixtures containing
benzene, toluene, and o-xylene at the 10-ppm level [160].
Manginell et al. reported a microfabricated planar preconcentrator for integration with a silicon-based gas chromatographic column [161]. This silicon-based device consisted
of a flow-through chamber formed by through-wafer etching
down to a silicon nitride membrane, onto which a thin-film
Pt heater had been previously patterned. The heater served
as the hotplate, and was covered by a porous sol-gel adsorbent layer. Preconcentration factors of between 100 and 500
were achieved for the analyte of interest, dimethyl methyl
phosphonate (DMMP), a nerve gas simulant. Heating rates
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
of 10 C/s were possible, which represented a thousandfold
improvement over conventional systems. At the same time,
power consumption, which was 100 mW for a typical desorption temperature of 200 C, was a thousandfold lower.
Increased temperatures are also beneficial in GC for
separating less volatile compounds (i.e., compounds that are
strongly retained on the stationary phase). Noh et al. recently
presented a gas chromatographic column micromachined in
parylene, a polymer exhibiting a low heat capacity and good
chemical inertness [162]. Parylene was deposited from the
gas phase onto a DRIE mold, followed by an intermediate
layer of Pt, and finally a second layer of parylene. A Pyrex
wafer was then coated with parylene, and bonded to the
coated Si wafer under high pressure and temperature. The
Si was then etched away, leaving a freestanding parylene
column attached to the Pyrex wafer. Gold was evaporated
onto the tops of the column to serve as a heater. The purpose
of the intermediate layer of Pt in the parylene walls was
twofold: namely, it reduced gas permeation through the walls
of the column, in addition to diffusing the heat uniformly
throughout the column. Simulation and experiment showed
that this polymer material showed much faster heating and
cooling than chip-based silicon-glass columns.
A unique micro liquid handling method was reported by
Handique et al., who used an integrated heater and RTD to
thermally generate air pressure inside a microfluidic channel
for pumping discrete droplets [163]. The hybrid device contained a lower Si chip with Pt electrodes formed by liftoff and
coated by oxide, and an upper glass chip with the microfluidic channels and air trapping chambers (see Fig. 15). The
chamber containing the heater and trapped air is connected
to the microfluidic channel via a small channel. It is through
this channel that heated, expanding air pushes its way from
the chamber into the liquid-containing channel, displacing a
plug of liquid before it. Droplets of nL volume could be precisely defined using this system at flow rates of 20 nL/s using
air heating rates of 6 C/s.
C. Pumping
As mentioned previously, EOF has been a popular means
of pumping liquids in microfluidic devices. In most cases,
this is still achieved by immersing wire electrodes into
open reservoirs at the ends of the channels. The resulting
electrolytically generated gas bubbles are easily removed, as
they simply drift to the surface of the liquids in the reservoir
and vanish into the air. Smaller, more compact devices could
be realized by integration of the pumping electrodes into the
microchannels themselves. However, this would result in
the formation and trapping of gas bubbles in the channels,
which would interrupt the applied electric field and halt the
EOF unless gas dissipation was facilitated in some way. To
avoid this problem, electrodes integrated into the reservoirs
themselves have been described by Figeys et al. [164].
After 30- m-deep channels and reservoirs had been etched
into the glass wafer, Au electrodes were vacuum-deposited
in the reservoirs. The electrode pads were connected by
an 80- m-wide metal line to contact pads, at which the
high voltage was applied. Jeong et al. also integrated Au
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
Fig. 15. (a) Schematic of the discrete drop pumping device.
(b)–(e) How the pump operates: (b) Liquid is introduced at the inlet,
and drawn into the device by capillary forces up to the hydrophobic
patch (channel surface is not conducive to capillary flow). Air is
trapped in the chamber during this process. (c) The heater is turned
on, and the air inside the chamber is heated to the point that it
exceeds a threshold pressure and enters the liquid channel to push a
well-defined liquid droplet past the hydrophobic patch. (d) Further
heating leads to continued expansion of air, and further motion of
the liquid droplet as a result. (e) The air pressure is vented as the
droplet passes the vent channel. (Reprinted with permission from
[163]. Copyright 2001 American Chemical Society.)
electrodes into their Si–glass device for synchronized cyclic
CE [22]. McKnight et al. photolithographically patterned
and etched a Au–Ti layer on glass to form electrodes
[86]. A channel in PDMS was positioned perpendicular
to these electrodes, which extended across the channel. In
this case, gas dissipation through the gas-permeable PDMS
layer ensured that the electric field was not interrupted. It
was possible to generate EOF in just a short segment of a
microchannel in this way, causing solution to be hydraulically
displaced on either side of the pair of electrodes.
In fact, bubbles generated by electrolysis may be used
beneficially in a microfluidic device for pumping purposes.
Böhm et al. described a microfluidic device comprising a
947
channel/reservoir structure in silicon, capped by a Pyrex
glass cover onto which a set of Pt electrodes had been
fabricated [165], [166]. The interdigitated electrodes served
a dual purpose, since the bubbles were generated and their
volume simultaneously measured by impedance at these
electrodes. The bubbles displaced liquid in reservoir channels connected to the gas generation chamber as they grew,
which in turn dispensed the liquids into a third channel for
further use upstream. Flow rates in the order of nL/s were
possible with this system [165], [166].
IV. CONCLUSION
Clearly, micromachining technologies have played a crucial role in the development of microfluidics from the very
beginning. The realization of sub- L flow systems with features precise enough to achieve the desired performance for
most applications is difficult if not impossible without a photolithographic process involved somewhere in device fabrication. Hence, micromachined channel devices have become inherent to the microfluidics field. The association of
microfluidics with MEMS has extended beyond the fabrication of simple microchannels, however, as has been discussed. The MEMS field has utilized the ability to work with
different materials to construct devices with an increasing
number of functions. The microfluidics world is learning to
do the same.
One area of increasing interest is the integration of optical
elements into microfluidic devices, a subject which was
not dealt with in this paper. Both early and more recent
papers have focused on small-volume, long-pathlength
detector cells for application to absorbance detection [27],
[32], [33], [44], [51], [167]. Other groups have investigated
the direct microfabrication into microfluidic devices of
microlenses [168]–[171], integrated waveguides [25], [172],
and photodiodes [139], [173]–[176]. A dc microplasma on a
chip has also been reported as an optical emission detector
for GC [177]–[179]. Besides the possibility of smaller,
more portable devices, the integrated approach also has the
potential to alleviate problems associated with the alignment
of external detection elements with the microfluidic chip.
Increased sensitivity may also result if the number of optical
interfaces can be minimized through integration.
The advantages of microchip electrophoresis especially
for nucleic acid analysis have been widely touted. Generally, the increase in speed reported is on the order of
tenfold to a hundredfold relative to capillaries and slab-gel
formats, respectively (see, e.g., [180], [181]). Resolution
of peaks in the electropherogram obtained on the chip,
though sometimes lower than that observed in fused-silica
capillaries [181], [182], is comparable in all cases to the
equivalent experiments done in conventional CE systems.
Throughput can be increased even further by resorting
to multicapillary arrays in glass, as has been put forth
particularly by the Mathies group [149], [183]–[187], but
also described by Sassi et al. [188] and Huang et al. [189].
The most recent example from the Mathies group describes
a 384-lane capillary array electrophoresis bioanalyzer for
ultrahigh-throughput genetic analysis [190]. This device
948
was fabricated on a 20-mm-diameter glass wafer, and had
384 sample reservoirs and 96 common buffer reservoirs.
Fabrication of such a system represents a challenge, as all
384 lanes need to be exactly the same. Moreover, getting
solutions onto the device requires some form of automated
sample addition step. Improving the interface between the
microfluidic device and the macroscopic world for this
and many other applications is an area in which MEMS
technologies could provide some advances in the future.
Last but not least, the trend toward the integration of
more sample pretreatment onto microfluidic devices is now
established, particularly in the area of genetic analysis.
A number of researchers have reported the (almost) total
integration of nucleic acid analysis, including sample
pretreatment, onto polymer-based [191]–[193], glass-based
[194], and silicon-based [23], [24] devices. Of these, the
devices emerging from the Burns group [23], [24] are the
most sophisticated from a MEMS point of view, incorporating micromachined elements for micro liquid handling,
thermal reaction, electrophoretic separation, and detection
onto a single device. With batch-type processing, it may be
possible to scale out these types to large arrays for highly
parallel analysis. However, control of liquids in multibranch
systems will ultimately require the development of new
valving concepts. This is because present valveless concepts
based on electroosmotic flow or hydrophobic patches, while
appropriate for many simpler applications, are somewhat
limited due to their tendency toward leakiness. Microfabricated valves based on pneumatically actuated elastomer
membranes have recently been proposed for array applications in biochemistry [195], [196]. Thorsen et al. have
extended this concept to realize high-density microfluidic
chips with thousands of valves and hundreds of individually
addressable microchambers [197]. These truly impressive
devices, designed to be analogous to electronic ICs, prove
the feasibility of very large-scale valving in complex microfluidic systems. Here too, MEMS technologies will play
an important role in the future development of microfluidics.
ACKNOWLEDGMENT
The authors would like to thank all the members, past
and present, of the TAS team in the Sensors, Actuators,
and Microsystems Laboratory (SAMLAB), at the Institute
of Microtechnology, University of Neuchâtel, Neuchâtel,
Switzerland, for their valuable efforts. Their contributions to
this paper in the form of photos and figures are of particular
note. The authors would also like to thank G. C. Fiaccabrino
(SAMLAB), R. Guijt-van Duijn (Technical University of
Delft), T. Fujii (University of Tokyo), H. Minhas (RSC
Journals), and J. Rossier (Diagnoswiss S.A., Monthey,
Switzerland) for photos and figures.
REFERENCES
[1] A. Manz, N. Graber, and H. M. Widmer, “Miniaturized total chemical analysis systems: A novel concept for chemical sensing,” Sens.
Actuators, vol. B1, pp. 244–248, 1990.
[2] H. M. Widmer, “Trends in industrial analytical chemistry,” Trends
Anal. Chem., vol. 2, pp. viii–x, 1983.
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
[3] A. Manz, D. J. Harrison, E. M. J. Verpoorte, J. C. Fettinger, A.
Paulus, H. Ludi, and H. M. Widmer, “Planar chips technology
for miniaturization and integration of separation techniques into
monitoring systems—Capillary electrophoresis on a chip,” J.
Chromatogr., vol. 593, pp. 253–258, 1992.
[4] S. C. Jacobson, R. Hergenröder, L. B. Koutny, and J. M. Ramsey,
“High-speed separations on a microchip,” Anal. Chem., vol. 66, pp.
1114–1118, 1994.
[5] D. J. Harrison, K. Fluri, K. Seiler, Z. H. Fan, C. S. Effenhauser, and
A. Manz, “Micromachining a miniaturized capillary electrophoresisbased chemical analysis system on a chip,” Science, vol. 261, pp.
895–897, 1993.
[6] A. Manz, C. S. Effenhauser, N. Burggraf, D. J. Harrison, K. Seiler,
and K. Fluri, “Electroosmotic pumping and electrophoretic separation for miniaturised chemical analysis systems,” J. Micromech. Microeng., vol. 4, pp. 257–265, 1994.
[7] F. von Heeren, E. Verpoorte, A. Manz, and W. Thormann, “Micellar
electrokinetic chromatography separations and analyses of biological samples on a cyclic planar microstructure,” Anal. Chem., vol.
68, pp. 2044–2053, 1996.
[8] J. B. Knight, A. Vishwanath, J. P. Brody, and R. H. Austin, “Hydrodynamic focusing on a silicon chip: Mixing nanoliters in microseconds,” Phys. Rev. Lett., vol. 80, pp. 3863–3866, 1998.
[9] W. T. Ko and J. T. Suminto, “Semiconductor integrated circuit technology and micromachining,” in Sensors, W. Göpel, J. Hasse, and
J. N. Zemel, Eds. Weinheim, Germany: VCH, 1989, vol. 1, pp.
107–168.
[10] G. T. A. Kovacs, K. Petersen, and M. Albin, “Silicon micromachining—Sensors to systems,” Anal. Chem., vol. 68, pp.
A407–A412, 1996.
[11] G. T. A. Kovacs, Micromachined Transducers Sourcebook. New
York: McGraw-Hill, 1998.
[12] M. Madou, Fundamentals of Microfabrication, 1st ed. Boca
Raton, FL: CRC Press, 1997.
[13] K. E. Petersen, “Silicon as a mechanical material,” Proc. IEEE, vol.
70, pp. 420–457, May 1982.
[14] E. H. Klaassen, K. Petersen, J. M. Noworoski, J. Logan, N. I. Maluf,
J. Brown, C. Storment, W. McCulley, and G. T. A. Kovacs, “Silicon fusion bonding and deep reactive ion etching; a new technology
for microstructures,” in Dig. Tech. Papers Transducers’95, vol. 1, S.
Middelhoek and K. Cammann, Eds., 1995, pp. 556–559.
[15] J. Bhardwaj, H. Ashraf, and A. McQuarrie, “Dry silicon etching for
MEMS,” in Proc. 191st Meeting Electrochemical Society: Symposium III Microstructures and Microfabricated Systems, vol. 97-5,
1997, pp. 118–130.
[16] P. A. Clerc, L. Dellmann, F. Grétillat, M. A. Grétillat, P. F. Indermühle, S. Jeanneret, P. Luginbuhl, C. Marxer, T. L. Pfeffer, G. A.
Racine, S. Roth, U. Staufer, C. Stebler, P. Thiébaud, and N. F. de
Rooij, “Advanced deep reactive ion etching: A versatile tool for microelectromechanical systems,” J. Micromech. Microeng., vol. 8, pp.
272–278, 1998.
[17] S. C. Terry, J. H. Jerman, and J. B. Angel, “A gas chromatographic
air analyzer fabricated on a silicon wafer,” IEEE Trans. Electron Devices, vol. ED-26, pp. 1880–1887, Dec. 1979.
[18] D. J. Harrison, P. G. Glavina, and A. Manz, “Toward miniaturized
electrophoresis and chemical analysis systems on silicon—An alternative to chemical sensors,” Sens. Actuators B, Chem., vol. 10, pp.
107–116, 1993.
[19] S. J. Pace, “Silicon semiconductor wafer for analyzing micronic biological samples,” U.S. Patent 4 908 112, 1990.
[20] D. E. Raymond, A. Manz, and H. M. Widmer, “Continuous sample
pretreatment using a free-flow electrophoresis device integrated onto
a silicon chip,” Anal. Chem., vol. 66, pp. 2858–2865, 1994.
, “Continuous separation of high molecular weight compounds
[21]
using a microliter volume free-flow electrophoresis microstructure,”
Anal. Chem., vol. 68, pp. 2515–2522, 1996.
[22] Y. Jeong, S. Kim, K. Chun, J. Chang, and D. S. Chung, “Methodology for miniaturized CE and insulation on a silicon substrate,” Lab
Chip, vol. 1, pp. 143–147, 2001.
[23] 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,
and D. T. Burke, “Microfabricated structures for integrated DNA
analysis,” Proc. Nat. Acad. Sci., vol. 93, pp. 5556–5561, 1996.
[24] M. A. Burns, B. 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, and D. T. Burke, “An integrated
nanoliter DNA analysis device,” Science, vol. 282, pp. 484–487,
1998.
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
[25] N. J. Petersen, K. B. Mogensen, and J. P. Kutter, “Performance of an
in-plane detection cell with integrated waveguides for UV/Vis absorbance measurements on microfluidic separation devices,” Electrophoresis, vol. 23, pp. 3528–3536, 2002.
[26] A. Manz, Y. Miyahara, J. Miura, Y. Watanabe, H. Miyagi, and K.
Sato, “Design of an open-tubular column liquid chromatograph
using silicon chip technology,” Sens. Actuators B, Chem., vol. 1,
pp. 249–255, 1990.
[27] G. Ocvirk, E. Verpoorte, A. Manz, M. Grasserbauer, and H. M.
Widmer, “High performance liquid chromatography partially
integrated onto a silicon chip,” Anal. Methods Instrum., vol. 2, pp.
74–82, 1995.
[28] S. Cowen and D. H. Craston, “An on-chip miniature liquid chromatography system: Design, construction and characterization,” in
Micro Total Analysis Systems 1994: Proceedings of the TAS 1994
Symposium, A. van den Berg and P. Bergveld, Eds. Dordrecht, The
Netherlands: Kluwer, pp. 295–298.
[29] J. P. Murrihy, M. C. Breadmore, A. M. Tan, M. McEnery, J. Alderman, C. O’Mathuna, A. P. O’Neill, P. O’Brien, N. Advoldvic,
P. R. Haddad, and J. D. Glennon, “Ion chromatography on-chip,” J.
Chromatogr. A, vol. 924, pp. 233–238, 2001.
[30] A. P. O’Neill, P. O’Brien, J. Alderman, D. Hoffman, M. McEnery, J.
Murrihy, and J. D. Glennon, “On-chip definition of picolitre sample
injection plugs for miniaturised liquid chromatography,” J. Chromatogr. A, vol. 924, pp. 259–263, 2001.
[31] M. McEnery, A. M. Tan, J. Alderman, J. Patterson, S. C. O’Mathuna,
and J. D. Glennon, “Liquid chromatography on-chip: Progression
towards a micro total analysis system,” Analyst, vol. 125, pp. 25–27,
2000.
[32] E. Verpoorte, A. Manz, H. Ludi, A. E. Bruno, F. Maystre, B. Krattiger, H. M. Widmer, B. H. van der Schoot, and N. F. de Rooij, “A
silicon flow cell for optical detection in miniaturized total chemical
analysis systems,” Sens. Actuators B, Chem., vol. 6, pp. 66–70, 1992.
[33] R. M. Tiggelaar, T. T. Veenstra, R. G. P. Sanders, J. G. E. Gardeniers,
M. C. Elwenspoek, and A. van den Berg, “A light detection cell to
be used in a micro analysis system for ammonia,” Talanta, vol. 56,
pp. 331–339, 2002.
[34] M. W. Losey, R. J. Jackman, S. L. Firebaugh, M. A. Schmidt, and
K. F. Jensen, “Design and fabrication of microfluidic devices for
multiphase mixing and reaction,” J. Microelectromech. Syst., vol. 11,
pp. 709–717, 2002.
[35] M. W. Losey, M. A. Schmidt, and K. F. Jensen, “Microfabricated
multiphase packed-bed reactors: Characterization of mass transfer
and reactions,” Ind. Eng. Chem. Res., vol. 40, pp. 2555–2562, 2001.
[36] A. J. Franz, K. F. Jensen, and M. A. Schmidt, “Palladium-based
micromembranes for hydrogen separation and hydrogenation/dehydrogenation reactions,” in Microreaction Technology: Industrial
Prospects, W. Ehrfeld, Ed. Berlin, Germany: Springer-Verlag,
2000, pp. 267–276.
[37] R. Srinivasan, I. M. Hsing, P. E. Berger, K. F. Jensen, S. L. Firebaugh, M. A. Schmidt, M. P. Harold, J. J. Lerou, and J. F. Ryley,
“Micromachined reactors for catalytic partial oxidation reactions,”
Amer. Inst. Chem. Eng. J., vol. 43, pp. 3059–3069, 1997.
[38] E. M. J. Verpoorte, B. H. van der Schoot, S. Jeanneret, A. Manz,
H. M. Widmer, and N. F. de Rooij, “Three-dimensional micro
flow manifolds for minaturized chemical analysis systems,” J.
Micromech. Microeng., vol. 4, pp. 246–256, 1994.
[39] T. T. Veenstra, R. M. Tiggelaar, R. P. G. Sanders, J. W. Berenschot,
J. G. E. Gardeniers, J. M. Wissink, R. Mateman, M. C. Elwenspoek,
and A. van den Berg, “Monolithic versus modular integration of a
micro-FIA system for ammonium determination,” in Micro Total
Analysis Systems 2001: Proceedings of the TAS 2001 Symposium,
J. M. Ramsey and A. van den Berg, Eds. Dordrecht, The Netherlands: Kluwer, 2001, pp. 664–666.
[40] E. S. Kolesar and R. R. Reston, “Silicon-micromachined gas chromatography system used to separate and detect ammonia and nitrogen-dioxide. 2. Evaluation, analysis, and theoretical modeling of
the gas chromatography system,” J. Microelectromech. Syst., vol. 3,
pp. 147–154, 1994.
[41] R. R. Reston and E. S. Kolesar, “Silicon-micromachined gas
chromatography system used to separate and detect ammonia and
nitrogen-dioxide. 1. Design, fabrication, and integration of the gas
chromatography system,” J. Microelectromech. Syst., vol. 3, pp.
134–146, 1994.
[42] U. Lehmann, O. Krusemark, and J. Muller, “Micromachined gas
chromatograph based on a plasma polymerised stationary phase,”
in Micro Total Analysis Systems 2000: Proceedings of the TAS
2000 Symposium, A. van den Berg, W. Olthuis, and P. Bergveld,
Eds. Dordrecht, The Netherlands: Kluwer, 2000, pp. 167–170.
949
[43] G. C. Frye-Mason, R. J. Kottenstette, C. D. Mowry, C. H. Morgan,
R. P. Manginell, P. R. Lewis, C. M. Matzke, G. R. Dulleck, L.
F. Anderson, and D. R. Adkins, “Expanding the capabilities and
application of gas phase miniature chemical analysis systems
(ChemLab),” in Micro Total Analysis Systems 2001: Proceedings
of the TAS 2001 Symposium, J. M. Ramsey and A. van den Berg,
Eds. Dordrecht, The Netherlands: Kluwer, 2001, pp. 658–660.
[44] A. Daridon, M. Sequeira, G. Pennarun-Thomas, H. Dirac, J. P. Krog,
P. Gravesen, J. Lichtenberg, D. Diamond, E. Verpoorte, and N. F. de
Rooij, “Chemical sensing using an integrated microfluidic system
based on the Berthelot reaction,” Sens. Actuators B, Chem., vol. 76,
pp. 235–243, 2001.
[45] D. Juncker, H. Schmid, U. Drechsler, H. Wolf, M. Wolf, B. Michel,
N. de Rooij, and E. Delamarche, “Autonomous microfluidic capillary system,” Anal. Chem., vol. 74, pp. 6139–6144, 2002.
[46] L. A. Christel, K. Petersen, W. McMillan, and M. A. Northrup,
“Rapid, automated nucleic acid probe assays using silicon microstructures for nucleic acid concentration,” J. Biomech. Eng., vol.
121, pp. 22–27, 1999.
[47] P. Thiébaud, C. Beuret, M. Koudelka-Hep, M. Bove, S. Martinoia,
M. Grattarola, H. Jahnsen, R. Rebaudo, M. Balestrino, J. Zimmer,
and Y. Dupont, “An array of Pt-tip microelectrodes for extracellular
monitoring of activity of brain slices,” Biosens. Bioelectron., vol. 14,
pp. 61–65, 1999.
[48] M. A. Grétillat, F. Paoletti, P. Thiébaud, S. Roth, M. Koudelka-Hep,
and N. F. de Rooij, “A new fabrication method for borosilicate glass
capillary tubes with lateral inlets and outlets,” Sens. Actuators A,
Phys., vol. 60, pp. 219–222, 1997.
[49] A. Dodge, K. Fluri, E. Verpoorte, and N. F. de Rooij, “Electrokinetically driven microfluidic chips with surface modified chambers
for heterogeneous immunoassays,” Anal. Chem., vol. 73, pp.
3400–3409, 2001.
[50] J. Lichtenberg, E. Verpoorte, and N. F. de Rooij, “Sample preconcentration by field amplification stacking for microchip-based capillary
electrophoresis,” Electrophoresis, vol. 22, pp. 258–271, 2001.
[51] A. Daridon, V. Fascio, J. Lichtenberg, R. Wütrich, H. Langen, E.
Verpoorte, and N. F. de Rooij, “Multi-layer microfluidic glass chips
for microanalytical applications,” Fresenius J. Anal. Chem., vol. 371,
pp. 261–269, 2001.
[52] S. J. Haswell, “Development and operating characteristics of micro
flow injection analysis systems based on electroosmotic flow—A
review,” Analyst, vol. 122, pp. R1–R10, 1997.
[53] T. McCreedy, “Fabrication techniques and materials commonly used
for the production of microreactors and micro total analytical systems,” Trends Anal. Chem., vol. 19, pp. 396–401, 2000.
[54] Z. H. Fan and D. J. Harrison, “Micromachining of capillary electrophoresis injectors and separators on glass chips and evaluation of
flow at capillary intersections,” Anal. Chem., vol. 66, pp. 177–184,
1994.
[55] S. C. Jacobson, R. Hergenröder, L. B. Koutny, R. J. Warmack, and
J. M. Ramsey, “Effects of injection schemes and column geometry
on the performance of microchip electrophoresis devices,” Anal.
Chem., vol. 66, pp. 1107–1113, 1994.
[56] P. C. Simpson, A. T. Woolley, and R. A. Mathies, “Microfabrication technology for the production of capillary array electrophoresis
chips,” Biomed. Microdevices, vol. 1, pp. 7–26, 1998.
[57] D. Sobek, S. D. Senturia, and M. L. Gray, “Microfabricated fused
silica flow chambers for flow cytometry,” in Proc. IEEE Solid-State
Sens. Actuator Workshop, 1994, pp. 260–263.
[58] S. C. Jacobson, A. W. Moore, and J. M. Ramsey, “Fused quartz
substrates for microchip electrophoresis,” Anal. Chem., vol. 67, pp.
2059–2063, 1995.
[59] K. Fluri, G. Fitzpatrick, N. Chiem, and D. J. Harrison, “Integrated
capillary electrophoresis devices with an efficient postcolumn reactor in planar quartz and glass chips,” Anal. Chem., vol. 68, pp.
4285–4290, 1996.
[60] R. M. Guijt, E. Baltussen, G. van der Steen, J. Frank, H. A. H. Billiet, T. Schalkhammer, F. Laugere, M. J. Vellekoop, A. Berthold,
P. M. Sarro, and G. W. K. van Dedem, “Capillary electrophoresis
with on-chip four-electrode capacitively coupled conductivity detection for application in bioanalysis,” Electrophoresis, vol. 22, pp.
2537–2541, 2001.
[61] H. Nakanishi, T. Nishimoto, R. Nakamura, A. Yotsumoto, T.
Yoshida, and S. Shoji, “Studies on SiO -SiO bonding with hydrofluoric acid. Room temperature and low stress bonding technique
for MEMS,” Sens. Actuators A, Phys., vol. 79, pp. 237–244, 2000.
[62] H. Y. Wang, R. S. Foote, S. C. Jacobson, J. H. Schneibel, and J.
M. Ramsey, “Low temperature bonding for microfabrication of
chemical analysis devices,” Sens. Actuators B, Chem., vol. 45, pp.
199–207, 1997.
950
[63] T. Ito, K. Sobue, and S. Ohya, “Water glass bonding for micro-total
analysis system,” Sens. Actuators B, Chem., vol. 81, pp. 187–195,
2002.
[64] N. Chiem, L. Lockyear-Shultz, P. Andersson, C. Skinner, and D.
J. Harrison, “Room temperature bonding of micromachined glass
devices for capillary electrophoresis,” Sens. Actuators B, Chem., vol.
63, pp. 147–152, 2000.
[65] A. Homsy, J. Lichtenberg, C. Massin, F. Vincent, P.-A. Besse,
R. S. Popovic, N. F. de Rooij, and E. Verpoorte, “Fabrication of
microfluidic channels with symmetric cross-sections for integrated
NMR analysis,” in Micro Total Analysis Systems 2002: Proceedings
of the TAS 2002 Symposium, A. van den Berg, Ed. Dordrecht,
The Netherlands: Kluwer, 2002, vol. 1, pp. 115–117.
[66] Y. Kikutani, T. Horiuchi, K. Uchiyama, H. Hisamoto, M. Tokeshi,
and T. Kitamori, “Glass microchip with three-dimensional microchannel network for 2 3 2 parallel synthesis,” Lab Chip, vol. 2,
pp. 188–192, 2002.
[67] Y. Kikutani, A. Hibara, K. Uchiyama, H. Hisamoto, M. Tokeshi,
and T. Kitamori, “Pile-up glass microreactor,” Lab Chip, vol. 2, pp.
193–196, 2002.
[68] B. He, N. Tait, and F. E. Regnier, “Fabrication of nanocolumns for
liquid chromatography,” Anal. Chem., vol. 70, pp. 3790–3797, 1998.
[69] B. He and F. E. Regnier, “Microfabricated liquid chromatography
columns based on collocated monolith support structures,” J. Pharmaceut. Biomed. Anal., vol. 17, pp. 925–932, 1998.
[70] B. He, J. Ji, and F. E. Regnier, “Capillary electrochromatography of
peptides in a microfabricated system,” J. Chromatogr. A, vol. 853,
pp. 257–262, 1999.
[71] X. Li, T. Abe, and M. Esashi, “Deep reactive ion etching of Pyrex
glass using SF plasma,” Sens. Actuators A, Phys., vol. 87, pp.
139–145, 2001.
[72] L. Ceriotti, K. Weible, N. F. de Rooij, and E. Verpoorte, “Rectangular
channels for lab-on-a-chip applications,” Microelectron. Eng., vol.
67–68, pp. 865–871, 2003, to be published.
[73] L. Ceriotti, E. Verpoorte, K. Weible, and N. F. De Rooij, “High aspect ratio channels for capillary electrophoresis,” in Digest of Technical Papers: Transducers 2001, E. Obermeier, Ed. Berlin, Germany: Springer-Verlag, 2001, vol. 2, pp. 1174–1177.
[74] T. R. Dietrich, W. Ehrfeld, M. Lacher, M. Kramer, and B. Speit,
“Fabrication technologies for microsystems utilizing photoetchable
glass,” Microelectron. Eng., vol. 30, pp. 497–504, 1996.
[75] A. Ruf, J. Diebel, M. Abraham, T. R. Dietrich, and M. Lacher,
“Ultra-long glass tips for atomic force microscopy,” J. Micromech.
Microeng., vol. 6, pp. 254–260, 1996.
[76] H. Becker, M. Arundell, A. Harnisch, and D. Hulsenberg, “Chemical
analysis in photostructurable glass chips,” Sens. Actuators B, Chem.,
vol. 86, pp. 271–279, 2002.
[77] Y. Fintschenko, P. Fowler, V. Spiering, G.-J. Burger, and A. van
den Berg, “Characterization of silicon-based insulated channels
for capillary electrophoresis,” in Micro Total Analysis Systems’98:
Proceedings of the TAS’98 Workshop, D. J. Harrison and A. van
den Berg, Eds. Dordrecht, The Netherlands: Kluwer, 1998, pp.
327–330.
[78] Y. Fintschenko and A. van den Berg, “Silicon microtechnology and
microstructures in separation science,” J. Chromatogr. A, vol. 819,
pp. 3–12, 1998.
[79] R. M. Guijt, E. Baltussen, G. van der Steen, R. B. M. Schasfoort, S.
Schlautmann, H. A.H. Billiet, J. Frank, G. W. van Dedem, and A.
van den Berg, “New approaches for fabrication of microfluidic capillary electrophoresis devices with on-chip conductivity detection,”
Electrophoresis, vol. 22, pp. 235–241, 2001.
[80] R. B. M. Schasfoort, S. Schlautmann, L. Hendrikse, and A. van
den Berg, “Field-effect flow control for microfabricated fluidic networks,” Science, vol. 286, pp. 942–945, 1999.
[81] Y. N. Xia and G. M. Whitesides, “Soft lithography,” Angew.
Chem.-Int. Edit. Engl., vol. 37, pp. 551–575, 1998.
[82] B. Michel, A. Bernard, A. Bietsch, E. Delamarche, M. Geissler,
D. Juncker, H. Kind, J. P. Renault, H. Rothuizen, H. Schmid, P.
Schmidt-Winkel, R. Stutz, and H. Wolf, “Printing meets lithography: Soft approaches to high-resolution patterning,” IBM J. Res.
Dev., vol. 45, pp. 697–719, 2001.
[83] J. C. McDonald, D. C. Duffy, J. R. Anderson, D. T. Chiu, H. Wu, O.
J. A. Schueller, and G. M. Whitesides, “Fabrication of microfluidic
systems in poly(dimethylsiloxane) [Review],” Electrophoresis, vol.
21, pp. 27–40, 2000.
[84] S. Masuda, M. Washizu, and T. Nanba, “Novel method of cell fusion
in field constriction area in fluid integrated circuit,” IEEE Trans. Ind.
Applicat., vol. 25, pp. 732–737, July/Aug. 1989.
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
[85] B. H. Jo, L. M. Van Lerberghe, K. M. Motsegood, and D. J. Beebe,
“Three-dimensional micro-channel fabrication in polydimethylsiloxane (PDMS) elastomer,” J. Microelectromech. Syst., vol. 9, pp.
76–81, 2000.
[86] T. E. McKnight, C. T. Culbertson, S. C. Jacobson, and J. M. Ramsey,
“Electroosmotically induced hydraulic pumping with integrated
electrodes on microfluidic devices,” Anal. Chem., vol. 73, pp.
4045–4049, 2001.
[87] D. C. Duffy, J. C. McDonald, O. J. A. Schueller, and G. M. Whitesides, “Rapid prototyping of microfluidic systems in poly(dimethylsiloxane),” Anal. Chem., vol. 70, pp. 4974–4984, 1998.
[88] L. Ceriotti, N. F. de Rooij, and E. Verpoorte, “An integrated fritless
column for on-chip capillary electrochromatography with conventional stationary phases,” Anal. Chem., vol. 74, pp. 639–647, 2002.
[89] C. S. Effenhauser, G. J. M. Bruin, A. Paulus, and M. Ehrat,
“Integrated capillary electrophoresis on flexible silicone microdevices—Analysis of DNA restriction fragments and detection of
single DNA molecules on microchips,” Anal. Chem., vol. 69, pp.
3451–3457, 1997.
[90] H. P. Chou, C. Spence, A. Scherer, and S. Quake, “A microfabricated
device for sizing and sorting DNA molecules,” Proc. Nat. Acad. Sci.,
vol. 96, pp. 11–13, 1999.
[91] G. Ocvirk, M. Munroe, T. Tang, R. Oleschuk, K. Westra, and
D. J. Harrison, “Electrokinetic control of fluid flow in native
poly(dimethylsiloxane) capillary electrophoresis devices,” Electrophoresis, vol. 21, pp. 107–115, 2000.
[92] V. Linder, E. Verpoorte, W. Thormann, N. F. de Rooij, and H.
Sigrist, “Surface biopassivation of replicated poly(dimethylsiloxane) microfluidic channels and application to heterogeneous
immunoreaction with on-chip fluorescence detection,” Anal. Chem.,
vol. 73, pp. 4181–4189, 2001.
[93] M. T. Gale, “Replication techniques for diffractive optical elements,”
Microelectron. Eng., vol. 34, pp. 321–339, 1997.
[94] J. W. Hong, T. Fujii, M. Seki, T. Yamamoto, and I. Endo, “Integration of gene amplification and capillary gel electrophoresis on
a poly(dimethylsiloxane)-glass hybrid microchip,” Electrophoresis,
vol. 22, pp. 328–333, 2001.
[95] J. R. Anderson, D. T. Chiu, R. J. Jackman, O. Cherniavskaya, J.
C. McDonald, H. K. Wu, S. H. Whitesides, and G. M. Whitesides,
“Fabrication of topologically complex three-dimensional microfluidic systems in PDMS by rapid prototyping,” Anal. Chem., vol. 72,
pp. 3158–3164, 2000.
[96] J. M. Shaw, J. D. Gelorme, N. C. LaBianca, W. E. Conley, and S. J.
Holmes, “Negative photoresists for optical lithography,” IBM J. Res.
Dev., vol. 41, pp. 81–94, 1997.
[97] H. Lorenz, M. Despont, N. Fahrni, N. LaBianca, P. Renaud, and
P. Vettiger, “SU-8: a low-cost negative resist for MEMS,” J. Micromech. Microeng., vol. 7, pp. 121–124, 1997.
[98] H. Lorenz, M. Despont, N. Fahrni, J. Brugger, P. Vettiger, and P.
Renaud, “High-aspect-ratio, ultrathick, negative-tone near-UV photoresist and its applications for MEMS,” Sens. Actuators A, Phys.,
vol. 64, pp. 33–39, 1998.
[99] O. Hofmann, P. Niedermann, and A. Manz, “Modular approach
to fabrication of three-dimensional microchannel systems in
PDMS—Application to sheath flow microchips,” Lab Chip, vol. 1,
pp. 108–114, 2001.
[100] E. Leclerc, Y. Sakai, and T. Fujii, “Human cancerous liver cells culture in a PDMS (PolyDiMethylSiloxane) network of micro channels
and chambers,” in Proc. IEEE-EMBS Special Topic Conf. Molecular,
Cellular and Tissue Engineering, 2002, pp. 104–105.
[101] H. Becker and C. Gärtner, “Polymer microfabrication methods for
microfluidic analytical applications,” Electrophoresis, vol. 21, pp.
12–26, 2000.
[102] H. Becker and L. E. Locascio, “Polymer microfluidic devices,” Talanta, vol. 56, pp. 267–287, 2002.
[103] L. Martynova, L. E. Locascio, M. Gaitan, G. W. Kramer, R.
G. Christensen, and W. A. MacCrehan, “Fabrication of plastic
microfluid channels by imprinting methods,” Anal. Chem., vol. 69,
pp. 4783–4789, 1997.
[104] J. Xu, L. Locascio, M. Gaitan, and C. S. Lee, “Room-temperature
imprinting method for plastic microchannel fabrication,” Anal.
Chem., vol. 72, pp. 1930–1933, 2000.
[105] R. M. McCormick, R. J. Nelson, M. G. Alonso-Amigo, D. J. Benvegnu, and H. H. Hooper, “Microchannel electrophoretic separations
of DNA in injection-molded plastic substrates,” Anal. Chem., vol.
69, pp. 2626–2630, 1997.
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
[106] N. Thomas, A. Ocklind, I. Blikstad, S. Griffiths, M. Kenrick, H. Derand, G. Ekstrand, C. Ellström, A. Larsson, and P. Andersson, “Integrated cell based assays in microfabricated disposable CD devices,”
in Micro Total Analysis Systems 2000: Proceedings of the TAS
2000 Symposium, A. van den Berg, W. Olthuis, and P. Bergveld,
Eds. Dordrecht, The Netherlands: Kluwer, 2000, pp. 249–252.
[107] G. J. Kellogg, T. E. Arnold, B. L. Carvalho, D. C. Duffy, and
N. F. Sheppard Jr., “Centrifugal microfluidics: Applications,” in
Micro Total Analysis Systems 2000: Proceedings of the TAS
2000 Symposium, A. van den Berg, W. Olthuis, and P. Bergveld,
Eds. Dordrecht, The Netherlands: Kluwer, 2000, pp. 239–242.
[108] C. Elsner, J. Dienelt, and D. Hirsch, “3D-microstructure replication
processes using UV-curable acrylates,” Microelectron. Eng., vol. 65,
pp. 163–170, 2003.
[109] M. A. Roberts, J. S. Rossier, P. Bercier, and H. Girault, “UV laser
machined polymer substrates for the development of microdiagnostic systems,” Anal. Chem., vol. 69, pp. 2035–2042, 1997.
[110] A. C. Henry, E. A. Waddell, R. Shreiner, and L. E. Locascio, “Control of electroosmotic flow in laser-ablated and chemically modified
hot imprinted poly(ethylene terephthalate glycol) microchannels,”
Electrophoresis, vol. 23, pp. 791–798, 2002.
[111] T. J. Johnson, D. Ross, M. Gaitan, and L. E. Locascio, “Laser modification of preformed polymer microchannels: Application to reduce
band broadening around turns subject to electrokinetic flow,” Anal.
Chem., vol. 73, pp. 3656–3661, 2001.
[112] T. J. Johnson, D. Ross, and L. E. Locascio, “Rapid microfluidic
mixing,” Anal. Chem., vol. 74, pp. 45–51, 2002.
[113] H. Klank, J. P. Kutter, and O. Geschke, “CO -laser micromachining
and back-end processing for rapid production of PMMA-based microfluidic systems,” Lab Chip, vol. 2, pp. 242–246, 2002.
[114] B. H. Weigl, R. Bardell, T. Schulte, F. Battrell, and J. Hayenga, “Design and rapid prototyping of thin-film laminate-based microfluidic
devices,” Biomed. Microdevices, vol. 3, pp. 267–274, 2001.
[115] J. S. Rossier, C. Vollet, A. Carnal, G. Lagger, V. Gobry, H. H. Girault,
P. Michel, and F. Reymond, “Plasma etched polymer microelectrochemical systems,” Lab Chip, vol. 2, pp. 145–150, 2002.
[116] M. A. Schwarz and P. C. Hauser, “Recent developments in detection
methods for microfabricated analytical devices,” Lab Chip, vol. 1,
pp. 1–6, 2001.
[117] N. A. Lacher, K. E. Garrison, R. S. Martin, and S. M. Lunte,
“Microchip capillary electrophoresis/electrochemistry,” Electrophoresis, vol. 22, pp. 2526–2536, 2001.
[118] C. Haber, “Electrochemical detection in capillary electrophoresis,”
in Handbook of Capillary Electrophoresis, 2nd ed, J. P. Landers,
Ed. Boca Raton, FL: CRC Press, 1997, pp. 425–447.
[119] A. Manz, D. J. Harrison, E. Verpoorte, and H. M. Widmer, “Planar
chips technology for miniaturization of separation systems: A developing perspective in chemical monitoring,” in Advances in Chromatography, P. R. Brown and E. Grushka, Eds. New York: Marcel
Dekker, 1993, vol. 33, pp. 1–65.
[120] A. J. Gawron, R. S. Martin, and S. M. Lunte, “Fabrication
and evaluation of a carbon-based dual-electrode detector for
poly(dimethylsiloxane) electrophoresis chips,” Electrophoresis,
vol. 22, pp. 242–248, 2001.
[121] R. S. Martin, K. L. Ratzlaff, B. H. Huynh, and S. M. Lunte,
“In-channel electrochemical detection for microchip capillary
electrophoresis using an electrically isolated potentiostat,” Anal.
Chem., vol. 74, pp. 1136–1143, 2002.
[122] R. Kikura-Hanajiri, R. S. Martin, and S. M. Lunte, “Indirect measurement of nitric oxide production by monitoring nitrate and nitrite
using microchip electrophoresis with electrochemical detection,”
Anal. Chem., vol. 74, pp. 6370–6377, 2002.
[123] M. Galloway and S. A. Soper, “Contact conductivity detection of
polymerase chain reaction products analyzed by reverse-phase ion
pair microcapillary electrochromatography,” Electrophoresis, vol.
23, pp. 3760–3768, 2002.
[124] M. Galloway, W. Stryjewski, A. Henry, S. M. Ford, S. Llopis, R.
L. McCarley, and S. A. Soper, “Contact conductivity detection in
poly(methyl methacrylate)-based microfluidic devices for analysis
of mono- and polyanionic molecules,” Anal. Chem., vol. 74, pp.
2407–2415, 2002.
[125] M. A. Schwarz, B. Galliker, K. Fluri, T. Kappes, and P. C. Hauser, “A
two-electrode configuration for simplified amperometric detection
in a microfabricated electrophoretic separation device,” Analyst, vol.
126, pp. 147–151, 2001.
951
[126] R. S. Martin, A. J. Gawron, B. A. Fogarty, F. B. Regan, E. Dempsey,
and S. M. Lunte, “Carbon paste-based electrochemical detectors for
microchip capillary electrophoresis/electrochemistry,” Analyst, vol.
126, pp. 277–280, 2001.
[127] J. S. Rossier, M. A. Roberts, R. Ferrigno, and H. H. Girault, “Electrochemical detection in polymer microchannels,” Anal. Chem., vol.
71, pp. 4294–4299, 1999.
[128] J. S. Rossier, A. Schwarz, F. Reymond, R. Ferrigno, F. Bianchi, and
H. H. Girault, “Microchannel networks for electrophoretic separations,” Electrophoresis, vol. 20, pp. 727–731, 1999.
[129] J. S. Rossier, R. Ferrigno, and H. H. Girault, “Electrophoresis with
electrochemical detection in a polymer microdevice,” J. Electroanal.
Chem., vol. 492, pp. 15–22, 2000.
[130] J. S. Rossier and H. H. Girault, “Enzyme linked immunosorbent
assay on a microchip with electrochemical detection,” Lab Chip, vol.
1, pp. 153–157, 2001.
[131] J. Wang, B. Tian, and E. Sahlin, “Integrated electrophoresis
chips/amperometric detection with sputtered gold working electrodes,” Anal. Chem., vol. 71, pp. 3901–3904, 1999.
[132] A. Hilmi and J. H. T. Luong, “Electrochemical detectors prepared
by electroless deposition for microfabricated electrophoresis chips,”
Anal. Chem., vol. 72, pp. 4677–4682, 2000.
[133] R. S. Martin, A. J. Gawron, S. M. Lunte, and C. S. Henry, “Dualelectrode electrochemical detection for poly(dimethylsiloxane)-fabricated capillary electrophoresis microchips,” Anal. Chem., vol. 72,
pp. 3196–3202, 2000.
[134] Y. Liu, J. C. Fanguy, J. M. Bledsoe, and C. S. Henry, “Dynamic
coating using polyelectrolyte multilayers for chemical control of
electroosmotic flow in capillary electrophoresis microchips,” Anal.
Chem., vol. 72, pp. 5939–5944, 2000.
[135] E. L’Hostis, P. E. Michel, G. C. Fiaccabrino, D. J. Strike, N. F. de
Rooij, and M. Koudelka-Hep, “Microreactor and electrochemical
detectors fabricated using Si and EPON SU-8,” Sens. Actuators B,
Chem., vol. 64, pp. 156–162, 2000.
[136] G. C. Fiaccabrino, X. M. Tang, N. Skinner, N. F. de Rooij, and
M. Koudelka-Hep, “Interdigitated microelectrode arrays based on
sputtered carbon thin-films,” Sens. Actuators B, Chem., vol. 35, pp.
247–254, 1996.
[137] A. T. Woolley, K. Lao, A. N. Glazer, and R. A. Mathies, “Capillary electrophoresis chips with integrated electrochemical detection,” Anal. Chem., vol. 70, pp. 684–688, 1998.
[138] D. J. Strike, G.-C. Fiaccabrino, M. Koudelka-Hep, and N. F. de
Rooij, “Enzymatic microreactor using Si, glass and EPON SU-8,”
Biomed. Microdevices, vol. 2, pp. 175–178, 2000.
[139] G. C. Fiaccabrino, N. F. de Rooij, and M. Koudelka-Hep, “On-chip
generation and detection of electrochemiluminescence,” Anal.
Chim. Acta., pp. 263–267, 1998.
[140] B. Timmer, W. Sparreboom, W. Olthuis, P. Bergveld, and A. van
den Berg, “Optimization of an electrolyte conductivity detector for
measuring low ion concentrations,” Lab Chip, vol. 2, pp. 121–124,
2002.
[141] A. Arora, J. C. T. Eijkel, W. E. Morf, and A. Manz, “A wireless
electrochemiluminescence detector applied to direct and indirect detection for electrophoresis on a microfabricated glass device,” Anal.
Chem., vol. 73, pp. 3282–3288, 2001.
[142] O. T. Guenat, W. E. Morf, B. H. van der Schoot, and N. F. de Rooij,
“Universal coulometric nanotitrators with potentiometric detection,”
Anal. Chim. Acta., pp. 261–272, 1998.
[143] O. T. Guenat, B. H. van der Schoot, W. E. Morf, and N. F. de Rooij,
“Triangle-programmed coulometric nanotitrations completed by
continuous flow with potentiometric detection,” Anal. Chem., vol.
72, pp. 1585–1590, 2000.
[144] R. P. Baldwin, T. J. Roussel Jr., M. M. Crain, V. Bathlagunda, D.
J. Jackson, J. Gullapalli, J. A. Conklin, R. Pai, J. F. Naber, K. M.
Walsh, and R. S. Keynton, “Fully integrated on-chip electrochemical
detection for capillary electrophoresis in a microfabricated device,”
Anal. Chem., vol. 74, pp. 3690–3697, 2002.
[145] J. Lichtenberg, N. F. de Rooij, and E. Verpoorte, “A microchip
electrophoresis system with integrated in-plane electrodes for
contactless conductivity detection,” Electrophoresis, vol. 23, pp.
3769–3780, 2002.
[146] K. F. Jensen, “Microreaction engineering—Is small better?,” Chem.
Eng. Sci., vol. 56, pp. 293–303, 2001.
[147] E. Verpoorte, “Microfluidic chips for clinical and forensic analysis,”
Electrophoresis, vol. 23, pp. 677–712, 2002.
952
[148] G. H. W. Sanders and A. Manz, “Chip-based microsystems for genomic and proteomic analysis,” Trends Anal. Chem., vol. 19, pp.
364–378, 2000.
[149] I. L. Medintz, B. M. Paegel, R. G. Blazej, C. A. Emrich, L. Berti, J.
R. Scherer, and R. A. Mathies, “High-performance genetic analysis
using microfabricated capillary array electrophoresis microplates,”
Electrophoresis, vol. 22, pp. 3845–3856, 2001.
[150] V. M. Ugaz, S. N. Brahmasandra, D. T. Burke, and M. A.
Burns, “Cross-linked polyacrylamide gel electrophoresis of
single-stranded DNA for microfabricated genomic analysis systems,” Electrophoresis, vol. 23, pp. 1450–1459, 2002.
[151] V. M. Ugaz, D. T. Burke, and M. A. Burns, “Microdevice-based
measurements of diffusion and dispersion in cross-linked and linear
polyacrylamide DNA sequencing gels,” Electrophoresis, vol. 23, pp.
2777–2787, 2002.
[152] T. Yamamoto, T. Nojima, and T. Fujii, “PDMS-glass hybrid microreactor array with embedded temperature control device. Application
to cell-free protein synthesis,” Lab Chip, vol. 2, pp. 197–202, 2002.
[153] A. Sanjoh and T. Tsukihara, “Spatiotemporal protein crystal growth
studies using microfluidic silicon devices,” J. Cryst. Growth, vol.
196, pp. 691–702, 1999.
[154] A. Sanjoh, T. Tsukihara, and S. Gorti, “Surface-potential controlled
Si-microarray devices for heterogeneous protein crystallization
screening,” J. Cryst. Growth, vol. 232, pp. 618–628, 2001.
[155] K. F. Jensen, I.-M. Hsing, R. Srinivasan, and M. A. Schmidt,
“Reaction engineering for microreactor systems,” in Microreactor
Technology: Proceedings of the First International Conference on
Microreaction Technology, W. Ehrfeld, Ed. Berlin, Germany:
Springer-Verlag, 1997, pp. 2–9.
[156] T. McCreedy and N. G. Wilson, “Microfabricated reactors for
on-chip heterogeneous catalysis,” Analyst, vol. 126, pp. 21–23,
2001.
[157] E. M. J. Verpoorte, B. H. van der Schoot, S. Jeanneret, A. Manz,
and N. F. de Rooij, “Silicon-based chemical microsensors and microsystems,” in Interfacial Design and Chemical Sensing, ser. ACS
Symposium Series. Washington, DC: American Chemical Society,
1994, vol. 561, pp. 244–254.
[158] J. C. T. Eijkel, A. Prak, S. Cowen, D. H. Craston, and A. Manz,
“Micromachined heated chemical reactor for pre-column derivatization,” J. Chromatogr. A, vol. 815, pp. 265–271, 1998.
[159] Y. Ueno, T. Horiuchi, T. Morimoto, and O. Niwa, “Microfluidic
device for airborne BTEX detection,” Anal. Chem., vol. 73, pp.
4688–4693, 2001.
[160] Y. Ueno, T. Horiuchi, M. Tomita, O. Niwa, H.-S. Zhou, T. Yamada,
and I. Honma, “Separate detection of BTX mixture gas by a microfluidic device using a function of nanosized pores of mesoporous
silica adsorbent,” Anal. Chem., vol. 74, pp. 5257–5262, 2002.
[161] R. P. Manginell, G. C. Frye-Mason, R. J. Kottenstette, P. R. Lewis,
and C. C. Wong, “Microfabricated planar preconcentrator,” in Tech.
Dig. Solid-State Sens. Actuator Workshop, 2000, pp. 179–182.
[162] H.-S. Noh, P. J. Hesketh, and G. C. Frye-Mason, “Parylene gas
chromatographic column for rapid thermal cycling,” J. Microelectromech. Syst., vol. 11, pp. 718–725, 2002.
[163] K. Handique, D. T. Burke, C. H. Mastrangelo, and M. A. Burns,
“On-chip thermopneumatic pressure for discrete drop pumping,”
Anal. Chem., vol. 73, pp. 1831–1838, 2001.
[164] D. Figeys, Y. B. Ning, and R. Aebersold, “A microfabricated device
for rapid protein identification by microelectrospray ion trap mass
spectrometry,” Anal. Chem., vol. 69, pp. 3153–3160, 1997.
[165] S. Böhm, B. Timmer, W. Olthuis, and P. Bergveld, “A closed-loop
controlled electrochemically actuated micro-dosing system,” J. Micromech. Microeng., vol. 10, pp. 498–504, 2000.
[166] S. Böhm, W. Olthuis, and P. Bergveld, “An integrated micromachined electrochemical pump and dosing system,” Biomed.
Microdevices, vol. 1, pp. 121–130, 1999.
[167] H. Salimi-Moosavi, Y. T. Jiang, L. Lester, G. McKinnon, and
D. J. Harrison, “A multireflection cell for enhanced absorbance
detection in microchip-based capillary electrophoresis devices,”
Electrophoresis, vol. 21, pp. 1291–1299, 2000.
[168] J.-C. Roulet, R. Völkel, H. P. Herzig, E. Verpoorte, N. F. de Rooij,
and R. Dändliker, “Microlens systems for fluorescence detection in
chemical microsystems,” Opt. Eng., vol. 40, pp. 814–821, 2001.
[169] J.-C. Roulet, R. Völkel, H. P. Herzig, E. Verpoorte, N. F. de Rooij,
and R. Dändliker, “Fabrication of multilayer systems combining microfluidic and microoptical elements for fluorescence detection,” J.
Microelectromech. Syst., vol. 10, pp. 482–491, 2001.
PROCEEDINGS OF THE IEEE, VOL. 91, NO. 6, JUNE 2003
[170]
[171]
[172]
[173]
[174]
[175]
[176]
[177]
[178]
[179]
[180]
[181]
[182]
[183]
[184]
[185]
[186]
[187]
[188]
, “Performance of an integrated micro-optical system for fluorescence detection in microfluidic systems,” Anal. Chem., vol. 74,
pp. 3400–3407, 2002.
J. Seo and L. P. Lee, “Integrated microfluidic optical systems
(iMOS) with LED,” in Micro Total Analysis Systems 2002: Proceedings of the TAS 2002 Symposium, Y. Baba, S. Shoji, and A.
van den Berg, Eds. Dordrecht, The Netherlands: Kluwer, 2002,
vol. 1, pp. 284–286.
P. Friis, K. Hoppe, O. Leistiko, K. B. Mogensen, J. Hübner, and
J. P. Kutter, “Monolithic integration of microfluidic channels and
optical waveguides in silica on silicon,” Appl. Optics, vol. 40, pp.
6246–6251, 2001.
P. E. Michel, G. C. Fiaccabrino, N. F. de Rooij, and M.
Koudelka-Hep, “Integrated sensor for continuous flow electrochemiluminescent measurements of codeine with different
ruthenium complexes,” Anal. Chim. Acta., vol. 392, pp. 95–103,
1999.
M. L. Chabinyc, D. T. Chiu, J. C. McDonald, A. D. Stroock, J. F.
Christian, A. M. Karger, and G. M. Whitesides, “An integrated fluorescence detection system in poly(dimethylsiloxane) for microfluidic applications,” Anal. Chem., vol. 73, pp. 4491–4498, 2001.
J. R. Webster, M. A. Burns, D. T. Burke, and C. H. Mastrangelo,
“Monolithic capillary electrophoresis device with integrated fluorescence detector,” Anal. Chem., vol. 73, pp. 1622–1626, 2001.
T. Kamei, J. R. Scherer, B. M. Paegel, A. M. Skelley, R. A.
Street, and R. A. Mathies, “Integrated amorphous silicon photodiode detector for microfabricated capillary electrophoresis
devices,” in Micro Total Analysis Systems 2002: Proceedings of
the TAS 2002 Symposium, Y. Baba, S. Shoji, and A. van den
Berg, Eds. Dordrecht, The Netherlands: Kluwer, 2002, vol. 1, pp.
257–259.
J. C. T. Eijkel, H. Stoeri, and A. Manz, “A molecular emission detector on a chip employing a direct current microplasma,” Anal.
Chem., vol. 71, pp. 2600–2606, 1999.
J. C. T. Eijkel, H. Stoeri, and A. Manz, “An atmospheric pressure
dc glow discharge on a microchip and its application as a molecular emission detector,” J. Anal. At. Spectrom., vol. 15, pp. 297–300,
2000.
, “A dc microplasma on a chip employed as an optical emission detector for gas chromatography,” Anal. Chem., vol. 72, pp.
2547–2552, 2000.
D. Schmalzing, L. Koutny, D. Chisholm, A. Adourian, P. Matsudaira, and D. Ehrlich, “Two-color multiplexed analysis of eight
short tandem repeat loci with an electrophoretic microdevice,”
Anal. Biochem., vol. 270, pp. 148–152, 1999.
N. J. Munro, K. Snow, J. A. Kant, and J. P. Landers, “Molecular diagnostics on microfabricated electrophoretic devices: From slab gelto capillary- to microchip-based assays for T- and B-cell lymphoproliferative disorders,” Clin. Chem., vol. 45, pp. 1906–1917, 1999.
H. Tian, A. Jaquins-Gerstl, N. Munro, M. Trucco, L. C. Brody, and J.
P. Landers, “Single-strand conformation polymorphism analysis by
capillary and microchip electrophoresis: A fast, simple method for
detection of common mutations in BRCA1 and BRCA2,” Genomics,
vol. 63, pp. 25–34, 2000.
P. C. Simpson, D. Roach, A. T. Woolley, T. Thorsen, R. Johnston, G.
F. Sensabaugh, and R. A. Mathies, “High-throughput genetic analysis using microfabricated 96-sample capillary array electrophoresis
microplates,” Proc. Nat. Acad. Sci., vol. 95, pp. 2256–2261, 1998.
Y. N. Shi, P. C. Simpson, J. R. Scherer, D. Wexler, C. Skibola, M.
T. Smith, and R. A. Mathies, “Radial capillary array electrophoresis
microplate and scanner for high-performance nucleic acid analysis,”
Anal. Chem., vol. 71, pp. 5354–5361, 1999.
I. Medintz, W. W. Wong, G. Sensabaugh, and R. A. Mathies,
“High speed single nucleotide polymorphism typing of a hereditary
haemochromatosis mutation with capillary array electrophoresis
microplates,” Electrophoresis, vol. 21, pp. 2352–2358, 2000.
I. L. Medintz, L. Berti, C. A. Emrich, J. Tom, J. R. Scherer, and
R. A. Mathies, “Genotyping energy-transfer-cassette-labeled shorttandem-repeat amplicons with capillary array electrophoresis microchannel plates,” Clin. Chem., vol. 47, pp. 1614–21, 2001.
B. M. Paegel, C. A. Emrich, G. J. Weyemayer, J. R. Scherer, and
R. A. Mathies, “High throughput DNA sequencing with a microfabricated 96-lane capillary array electrophoresis bioprocessor,” Proc.
Nat. Acad. Sci., vol. 99, pp. 574–579, 2002.
A. P. Sassi, A. Paulus, I. D. Cruzado, T. Bjornson, and H. H.
Hooper, “Rapid, parallel separations of D1S80 alleles in a plastic
microchannel chip,” J. Chromatogr. A, vol. 894, pp. 203–217, 2000.
VERPOORTE AND DE ROOIJ: MICROFLUIDICS MEETS MEMS
[189] Z. Huang, N. Munro, A. F. R. Huhmer, and J. P. Landers, “Acoustooptical deflection-based laser beam scanning for fluorescence detection on multichannel electrophoretic microchips,” Anal. Chem., vol.
71, pp. 5309–5314, 1999.
[190] C. A. Emrich, H. J. Tian, I. L. Medintz, and R. A. Mathies, “Microfabricated 384-lane capillary array electrophoresis bioanalyzer
for ultrahigh-throughput genetic analysis,” Anal. Chem., vol. 74, pp.
5076–5083, 2002.
[191] R. C. Anderson, X. Su, G. J. Bogdan, and J. Fenton. (2000)
A miniature integrated device for automated multistep genetic
assays. Nucleic Acids Res. [Online] Available: http://nar.oupjournals.org/cgi/content/full/28/12/e60
[192] P. Grodzinski, R. H. Liu, B. Chen, J. Blackwell, Y. Liu, D. Rhine, T.
Smekal, D. Ganser, C. Romero, H. Yu, T. Chan, and N. Kroutchinina,
“Development of plastic microfluidic devices for sample preparation,” Biomed. Microdevices, vol. 3, pp. 275–283, 2001.
[193] Y. Liu, C. B. Rauch, R. L. Stevens, R. Lenigk, J. Yang, D. B. Rhine,
and P. Grodzinski, “DNA amplification and hybridization assays
in integrated plastic monolithic devices,” Anal. Chem., vol. 74, pp.
3063–3070, 2002.
[194] B. M. Paegel, S. H. I. Yeung, and R. A. Mathies, “Microchip bioprocessor for integrated nanovolume sample purification and DNA
sequencing,” Anal. Chem., vol. 74, pp. 5092–5098, 2002.
[195] M. A. Unger, H. P. Chou, T. Thorsen, A. Scherer, and S. R. Quake,
“Monolithic microfabricated valves and pumps by multilayer soft
lithography,” Science, vol. 288, pp. 113–116, 2000.
[196] W. H. Grover, A. M. Skelley, C. N. Liu, E. T. Lagally, and R. A.
Mathies, “Practical valves and pumps for large-scale integration
into microfluidic analysis devices,” in Micro Total Analysis Systems
2002: Proceedings of the TAS 2002 Symposium, Y. Baba, S. Shoji,
and A. van den Berg, Eds. Dordrecht, The Netherlands: Kluwer,
2002, vol. 1, pp. 136–138.
[197] T. Thorsen, S. J. Maerkl, and S. R. Quake, “Microfluidic large-scale
integration,” Science, vol. 298, pp. 580–584, 2002.
Elisabeth Verpoorte received the Ph.D. in analytical chemistry from the
University of Alberta, Edmonton, AB, Canada, in 1990. Her dissertation
focused on the use of AC impedance analysis for studying the permeability
and behavior of dissociable species in PVC-based ion selective membranes.
She was in the Corporate Analytical Research department of Ciba Ltd.,
Basel, Switzerland, first in a postdoctoral position, and later in a permanent
position as Research Scientist. There, she worked on the development of
miniaturized total analysis systems (TAS). In July 1996, she assumed a
position as Team Leader in the group of Prof. N. F. de Rooij at the Institute
of Microtechnology (IMT), University of Neuchâtel, Switzerland, where her
research interests concentrated in the area of TAS (Lab-on-a-Chip) for bioanalytical and environmental applications. She led a research team of about
eight Ph.D. students and researchers while at IMT. Recently, she accepted
a Chair of Pharmaceutical Analysis at the Groningen Research Institute of
Pharmacy, University of Groningen, Groningen, The Netherlands. She holds
three patents and has published over 40 peer-reviewed papers.
Nico F. de Rooij (Fellow, IEEE) received the M.Sc. degree in physical
chemistry from the State University of Utrecht, Utrecht, The Netherlands,
in 1975, and the Ph.D. degree from Twente University of Technology,
Enschede, The Netherlands, in 1978.
From 1978 to 1982, he worked at the Research and Development Department of Cordis Europa NV, Roden, The Netherlands. In 1982, he joined
the Institute of Microtechnology of the University of Neuchâtel, Neuchâtel,
Switzerland (IMT UNI-NE), as Professor and Head of the Sensors, Actuators and Microsystems Laboratory. From 1990 to 1996 and since October
2002, he is Director of the IMT UNI-NE. He has lectured at the Swiss Federal Institute of Technology, Zurich (ETHZ), and since 1989, he has been a
part-time professor at the Swiss Federal Institute of Technology, Lausanne
(EPFL). He is a Member of the editorial boards for the journals Sensors and
Actuators and Sensors and Materials. His research activities include microfabricated sensors, actuators, and microsystems.
Dr. de Rooij was a Member of the steering committee of the International Conference on Solid-State Sensors and Actuators and of Eurosensors.
He acted as European Program Chairman of Transducers ’87 and General
Chairman of Transducers ’89. He is a Member of the editorial board of the
IEEE JOURNAL OF MICROELECTROMECHANICAL SYSTEMS.
953