3772
Swomitra K. Mohanty
Dongshin Kim
David J. Beebe
Department of Biomedical
Engineering,
University of Wisconsin-Madison,
Madison, WI, USA
Received April 17, 2006
Revised June 22, 2006
Accepted June 23, 2006
Electrophoresis 2006, 27, 3772–3778
Research Article
Do-it-yourself microelectrophoresis chips
with integrated sample recovery
We present a microelectrophoresis chip that is simple to fabricate using the mfluidic
tectonics (mFT) platform (Beebe, D. J. et al., Proc. Natl. Acad. Sci. USA 2000, 97,
13488–13493; Agarwal, A. K. et al., J. Micromech. Microeng. 2006, 16, 332–340). The
device contains a removable capillary insert (RCI) for easy sample collection after
separation (Atencia, J. et al., Lab Chip 2006, DOI: 10. 1039/b514068d). Device construction is accomplished in less than 20 min without specialized equipment traditionally associated with microelectrophoresis chip construction. mFT was used to build a
PAGE device utilizing two orthogonal microchannels. One channel performs standard
separations, while the second channel serves as an access point to remove bands of
interest from the chip via the RCI. The RCI contains an integrated electrode that facilitates the removal of bands using electrokinetic techniques. The device was characterized using prestained proteins (Pierce BlueRanger and TriChromRanger). Samples were loaded into the microelectrophoresis device via a standard micropipette. An
electrical field of 40 V/cm was used to separate and collect the proteins. The microPAGE device is simple to fabricate, benefits from microscale analysis, and includes an
on-chip collection scheme that interfaces the macroworld with the microworld.
Keywords: Microelectrophoresis / mfluidic tectonics / Removable capillary insert
DOI 10.1002/elps.200600238
1 Introduction
Miniaturization of electrophoresis analysis via microelectrophoresis chips has been an area of active research in
recent years [1–3]. Advantages of miniaturization include
reduced material usage, compact analysis space, and
high speed analysis. However obstacles to microelectrophoresis chips are cost and fabrication techniques.
These chips require expensive cleanroom facilities or
specialized equipment not available to most biological
researchers. In addition, most chips do not contain a
convenient method for sample collection, which may further discourage researchers from using the technology,
as sample analysis after separation is often needed.
Historically, electrophoresis is one of the most common
methods to separate charged molecules such as DNA
and proteins [4]. These molecules have unique physical
Correspondence: Mr. Swomitra Mohanty, Department of Biomedical Engineering, Room 2139 ECB, 1550 Engineering Drive, Madison,
WI 53706, USA
E-mail: [email protected]
Fax: 1608-265-9239
Abbreviations: ìFT, mfluidic tectonics; RCI, removable capillary
insert; PIBA, poly iso-bornyl acrylate
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
characteristics based on size, charge, and shape that
allow researchers to separate them in the presence of a
sieving matrix and electric field. Typical separations are
done in either slab gels or capillaries. Slab gel electrophoresis is a well-established technique that has been
used for decades. It has proven to be economical and
yield reliable analytical results. However, separations
done using gel electrophoresis are very time consuming
(often taking several hours to complete). In the 1990s CE
was introduced, which had several desirable attributes
including reduced separation time, reduced sample/
reagent use, and automated analysis. However, CE
requires expensive complex equipment and is not always
appropriate for all separations.
As microelectrophoresis chips are not readily available,
one alternative that some researchers have employed to
improve upon traditional methods is inexpensive microslab gel electrophoresis systems [5]. The microslab gels
made it possible to achieve higher electric field strength
with lower voltages, decreasing analysis time. However,
fabrication of microslab gels is cumbersome and creating small loading wells is difficult. Recently a simple
method for making microslab gels was reported [6]. This
method casts gels (25 6 30 6 0.6 mm) between a
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Miniaturization
Electrophoresis 2006, 27, 3772–3778
microscope glass slide (24 6 30 mm) and transparency
film. By cutting injection holes into the film, defined
injection wells in the gel were created by taking advantage of polymerization inhibition properties of oxygen.
The gel is polymerized at the film interface where oxygen
is not present. However, gel at the injection holes were
exposed to oxygen which inhibits polymerization. One
draw back to this method is that band collection after
separation is difficult. The size of the gel and space between the bands is very small making traditional band
cutting challenging.
Researchers continue to focus on improving electrophoresis through continued miniaturization [7–13]. As mentioned above, miniaturization improves electrophoretic
analysis by reducing material usage, analysis space, and
analysis time. Due to high surface to volume ratios at the
microscale, Joule heating and temperature gradients are
less problematic allowing for higher electric fields
strengths (compared to microslab gels) in shorter distances. In addition, the reduced space used for microelectrophoresis allows for development of high-throughput systems that can be interfaced with commercial
automation devices. One caveat to increased miniaturization is that sample collection after on-chip analysis
becomes increasingly difficult. The details for off chip
sample manipulations must be addressed for microelectrophoresis chips to be fully useful. In cases where
separation of an unknown DNA sample is conducted, a
specific band of interest is present that needs to be
extracted for further analysis (for example Southern blot,
PCR, DNA sequencing). The trend often observed in
microfluidics is that the more miniaturized analysis devices become, the more inclined these devices are to
use automated systems/custom instrumentation for
sample loading and recovery. However, these automated
liquid handling systems add to the cost of microelectrophoresis chips providing the same economical obsta-
3773
cle to researchers that CE does. Another issue in the
cost of microelectrophoresis chips is in the materials
used and fabrication methods. Using alternative materials and fabrication techniques can make microelectrophoresis chips more economical and accessible to life
science labs.
Various materials have been used to fabricate microelectrophoresis chips including polydimethylsiloxane (PDMS),
glass, and silicon [9, 14]. Each type of material provides
advantages and disadvantages with regard to ease of
fabrication, cost, and repeatable results (Table 1). PDMS
is a class of elastomers that is made up of an inorganic
siloxane backbone and organic methyl groups [15]. When
polymerized, PDMS is optically clear and generally considered non-toxic, making it a useful construction material for microfluidic devices involving biological material.
Using a process known as soft lithography [16], it is possible to make micro cell culture constructs [17], micro in
vitro fertilization devices [18], and electrophoresis devices
[14]. Although PDMS is easy to handle, it is not optimal for
electrophoretic separations without surface modification
[19]. For example the surface of PDMS attracts biomolecules causing them to stick to the channel instead of
separating out during electrophoresis. In addition, molding PDMS requires using relief master molds. The master
mold fabrication process can take several hours
depending on the thickness and feature size. Although a
master can be used many times to make a device, the
materials and equipment needed to make masters can be
costly.
Glass and silicon are materials that work well for microelectrophoresis devices utilizing acrylamide gel or capillary fluid because of their surface properties and precise
fabrication processes. Glass is the material used in traditional devices (PAGE electrophoresis systems), thus its
choice for microscale devices is logical. Acrylamide gel
polymerizes within glass, which helps create a uniform gel
Table 1. Comparison of microelectrophoresis fabrication methods
Fabrication technique
Advantages
Disadvantages
Silicon/glass microelectromechanical
system (MEMS)
Sub-micrometer resolution machining,
well-established techniques for repeatable
devices
Requires cleanroom and expensive specialized
equipment, time- and labor-intensive,
complicated device assembly
PDMS
10-mm resolution , simple fabrication
method, easy device assembly, selfbonding material
Additional surface modification needed for
electrophoresis separations, requires
fabrication of a relief master mold that adds
to material costs and construction time
mfluidic tectonics
Fast/simple fabrication method, no masters
needed, easy device assembly, no
bonding step required
Highest resolution 20–50 mm, limited to
photo-definable materials
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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S. K. Mohanty et al.
matrix for separation. For capillary separations, the glass
surface allows charged layers to develop facilitating EOF.
The charged layers are important for many types of
separations involving positive and negative charged molecules. Silicon is a material traditionally used in integrated
circuit chip fabrication. As a result, micromachining techniques have already been developed for silicon, making
microelectrophoresis chip fabrication achievable with little
process adjustment. Fabrication in silicon also allows for
integration of electronic components such as detectors,
heaters, and electrodes, making it very attractive for complete lab-on-a-chip applications. Since silicon fabrication
involves batch processing, lab-on-a-chip devices can be
made simultaneously on a massive scale reducing the cost
per chip. However, such mass produced chips are not
readily available to life science researchers. Successful
devices have been made using glass and silicon [20, 21].
However, they are not as simple to build as PDMS devices.
Traditional micromachining techniques are time consuming and require expensive cleanroom facilities not available
to most researchers. A typical glass electrophoresis device
takes several hours to complete. Assembly of devices is
also an issue generally requiring anodic bonding to create a
completely sealed channel. Anodic bonding requires extremely clean surfaces, relatively high temperatures, and
high voltages to achieve. It is more labor intensive than
PDMS bonding. While silicon and glass make excellent
microelectrophoresis chips, the particular fabrication
methodology that is traditionally employed makes it
impractical for most life sciences research laboratories.
We have addressed the issues of high cost microelectrophoresis chips and sample collection using alternative
materials and fabrication methods than those previously
described. We present a microelectrophoresis chip containing a removable sample collection component fabricated using the mfluidic tectonics (mFT) platform [22, 23].
The device is simple to fabricate with minimal costs and
equipment requirements. The mFT platform uses poly isobornyl acrylate (PIBA) and liquid-phase polymerization to
pattern microstructures such as microfluidic channels,
mixers and valves [24]. PIBA is a high glass transition
temperature (Tg 887C) polymer which is solid at room
temperature. It has excellent adhesion properties to various substrates, such as glass, plastic, and metal. PIBA
can be polymerized by liquid-phase photopolymerization,
and shows minimal change (5–10%) in volume after polymerization. Photopolymerization of PIBA is done quickly
with typical polymerization times ranging from 5 to 40 s.
After polymerization, PIBA is stiff and optically clear. Devices are fabricated in a matter of minutes allowing for
quick design improvements if needed. This contrasts with
traditional glass and PDMS fabrication where both methods take hours to complete a device. While PDMS de© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Electrophoresis 2006, 27, 3772–3778
vices require a master (inverse of the desired shape) for
fabrication, mFT creates the channels via direct photopolymerization of the final device material eliminating the
need for masters. One other attractive aspect of mFT is no
bonding is required for creating microchannels since they
are automatically sealed during the channel formation.
2 Materials and methods
2.1 Device overview
A scheme for the microelectrophoresis device and removable capillary insert (RCI) [25] is shown in Fig. 1. The device contains two orthogonal channels with RCI. Each
RCI contains integrated electrodes (platinum wire) for
electric field application and sample recovery. When a
band of interest is identified, it is directed down the
attached orthogonal channel by activating the electrodes
at the end of the channel. This method of on-chip sample
extraction has been previously reported by Burns et al.
[26], giving complete details modeling electric fields at the
intersection and design criteria. The sample recovery
methods described here expand on this work by adding
the RCI for easy off-chip removal.
Figure 1. Scheme of microelectrophoresis device. Two
orthogonal channels are shown with an RCI at each end.
Each RCI has an integrated electrode to work as an
anode or cathode during electrophoretic separation and
collection. Proteins are initially sent down a separation
channel. When a band of interest is identified and reaches
the intersection, it is directed down the access channel by
activating the electrodes in that channel and shutting off
electric field in the separation channel.
2.2 Device fabrication
Microelectrophoresis chips were fabricated with three
layers of materials: glass-PIBA-glass (Fig. 2). Two adhesive spacers of 0.5 cm width, 5.5 cm length, and 125 mm
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Electrophoresis 2006, 27, 3772–3778
Miniaturization
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Figure 2. Fabrication process
for microelectrophoresis chip.
Microelectrophoresis chips are
fabricated of three layers of
materials, glass-PIBA-glass. (a)
Two adhesive spacers are
placed on the edges of the bottom glass. The top slide (No. 1
cover slip) is drilled with 1-mm
holes to provide access to the
patterned channels and to create reservoirs for sample loading/collection. (b) The cover
glass is placed on the adhesive
forming a chamber. (c) PIBA is
introduced
using
capillary
action and a (d) photo-mask is
visually aligned over the substrate. The substrate is exposed
to UV light. (e) Any un-polymerized PIBA is vacuumremoved, leaving an empty
channel ready for acrylamide
gel. (f) The RCI is placed at the
output of the separation channel
and ends of the access channel.
Acrylamide gel solution is
placed into the channel through
the RCI. This allows the gel to
not only fill the channel but to
remain in the RCI itself for polymerization. Once filled, the
device is exposed to UV light
again. This results in microchannels and RCI filled with
acrylamide gel. (g) Platinum wire
is placed in the RCI directly in
the gel and acts as an electrode
for the separation process. The
input reservoir of the channel
also has a platinum electrode in
electrical contact to serve as the
cathode for the separation.
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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S. K. Mohanty et al.
thickness (Grace Bio-Labs Secure-Seal Adhesive SA-S1L) were placed on the edges of the bottom glass (Corning 50 mm 6 75 mm plain microslide). The top slide
(Fisher No. 1 premium cover glass 35650 mm) was drilled with 1-mm holes using a diamond tip drill bit (Eurotool
Diamond Bur fine/med grit 1 mm, 1500 rpm) to provide
access holes for the patterned channels and to create
reservoirs for sample loading/collection. Since the cover
glass was so thin, drilling took less than 10 s per hole with
minimal breakage (typically one in eight slides cracked). A
photo-definable PIBA solution was made as described by
Beebe et al. [22]. The cover glass was placed on the
adhesive forming a chamber. PIBA was introduced using
capillary action and a photomask printed on an acetate
sheet with a clear field pattern was visually aligned over
the substrate. Features patterned for this work ranged
from 500 mm to 1000 mm in width by 3 cm in length; however, features sizes down to 20–50 mm can be created.
The substrate was exposed to UV light (Acticure EFOS
A4000) at 6.7 mW/cm2 for 17 s. PIBA was cured by UV
light, therefore areas of the mask that blocked light form
the channel shape. The ends of the channel were aligned
with the holes on the cover glass. A vacuum pump was
placed over the holes using a vacuum cup interface
(Integrated Dispensing Solutions part no. 8880264) to
remove any un-polymerized PIBA leaving an empty
sealed channel ready for acrylamide gel. Channels were
cleaned out using deionized (DI) water to remove any
PIBA residue on the surface of the glass.
2.3 Device interface and RCI construction
Electrophoresis 2006, 27, 3772–3778
wich served as the cathode for the separation. The electrode was kept 2 cm away from the loading zone to minimize effects on the protein markers (bubbles generated
from electrolysis). A drop of 0.56 Tris-borate-EDTA buffer
(TBE) was placed over the reservoir once the separation
process started to insure electric field continuity.
2.4 Device operation
Characterization of the device was done performing
separations on prestained protein markers (Pierce BlueRanger and TriChromRanger). TriChromRanger/BlueRanger consists of a stabilized and lyophilized formulation of seven proteins, ranging from 16.5 K to 210 K and is
easily visualized in normal light. The marker solution (1 mL)
was loaded into the channel reservoir. An electric field
strength of 40 V/cm was applied (Thermo EC 6000P
electrophoresis power supply) for 15 min. Typically in
other microfluidic systems electrokinetic injection is used
to inject samples into a channel [27]. This setup is complex and requires extra work and equipment (extra power
supplies, controllers, programing) to manipulate electric
fields. To simplify sample loading and make it more
accessible to life science labs, we used standard methods via a micropipette for the experiments described
here. Experiments were visualized on a standard stereoscope (Olympus SZX12) using Leica DFC300 FX camera
for imaging.
3 Results and discussion
3.1 Band separation/collection
Connectors to interface with the device were fabricated
out of PDMS via a coring process (no masters were used).
PDMS was poured over an adhesive (Grace Bio-Labs
Secure-Seal Adhesive SA-S-1L) and cured. Small 1 cm
6 1 cm squares were cut and holes were bored through
creating an access port. A one-inch piece of capillary
tubing (Cole-Palmer PTFE no. 06417–31) was inserted
into the connector forming the RCI. The RCI was placed
at the output of the separation channel and access
channel (Fig. 2f). UV curable acrylamide solution was
prepared using 500 mL 25% acrylamide/bisacrylamide
(37.5:1), 250 mL 1.5 M Tris-HCl at pH 8.8, 2 mL riboflavin
(0.2%), 50 ml TEMED (10%) and 448 mL of DI water to
yield a final concentration of 12.5%. Acrylamide gel solution was placed into the channel though the RCI (Fig. 2f).
Once filled, the device was exposed to UV light for 10 min
at 25 mW/cm2. This resulted in microchannels and RCIs
with polymerized acrylamide gel. A platinum wire was
placed in the RCI and acted as an electrode for the
separation process (Fig. 2g). The input reservoir of the
separation channel was in contact with a platinum wire
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
All experiments were conducted using 12.5% acrylamide
gel looking at band separation and post separation sample collection using the RCI (Fig. 3). Figure 3a shows the
top view of the device during operation. Figure 3b shows
seven bands of TriChromRanger as it separates. The
bands shown are myosin (210 K), phosphorylase B
(110 K), BSA (80 K), ovalbumin (47 K), carbonic anhydrase (32 K), trypsin inhibitor (25 K), and lysozyme
(16.5 K). Figure 3c shows an intersection with the
separation and access channel. As the band approached
the intersection, the electrodes in the collection channel
were activated (electrodes in the separation channel were
shut off) driving the band into the access channel towards
the RCI for collection.
3.2 Band-broadening and leakage
When the band reaches the intersection, some broadening in the shape occurs as the band is no longer confined within the separation channel. In addition, band
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Electrophoresis 2006, 27, 3772–3778
leakage occurs into channels at the cross-section when it
is not collected (Fig. 3d). These results are similar to that
reported by Lin et al. [26] and more detail regarding this
can be found in their publication. If a band of interest
comes after a band that is not collected, most of the
leaked material was removed from the path by activating
the electrodes in the access channel and driving the
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3777
material in the direction of a “waste” RCI directly opposite
of the RCI used for sample extraction. This is done before
the band of interest reaches the intersection. Occasional
bubbles could show up in the gel during fabrication as
seen in Fig. 3c. The bubbles did not have any major
impact on the performance of the microelectrophoresis
chip.
Figure 3. Separation and collection of proteins with microelectrophoresis device. (a) Top view of a
device during use. (b) Image of TriChromRanger proteins as separation is occurring. (c) Orthogonal
intersection of separation channel and access channel. The band of interest approaches the intersection. Once it is in range of the access channel, the electrodes at the end of the access channel are
activated directing the protein to the RCI where it can be collected. (d) Band leakage into the channel
was observed if a band was not collected. (e) The RCI with integrated electrode inserted into a PDMS
connector to interface with the device. (f) Acrylamide gel removed from the RCI after collecting the
protein. The protein is stored in a cylindrical shaped gel. Dark portion of the gel is BlueRanger protein.
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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S. K. Mohanty et al.
Electrophoresis 2006, 27, 3772–3778
3.3 Sample removal off-chip
5 References
Figure 3e shows an image of the RCI with the electrode
coming out of the capillary tubing. The RCI is inserted into
a PDMS connector to interface with the device. As mentioned above, acrylamide gel was polymerized in the RCI
and a positive potential was applied at the electrode to
drive the protein bands towards the RCI. After the sample
was collected inside the tubing, it was removed from the
chip by pulling the capillary out. The RCI provided a convenient package to transport the protein. The gel was
removed from the capillary by applying pressure at one
end leaving a cylindrical shaped gel. Figure 3f shows a
protein (dark color) filled gel from a capillary lying on a
microelectrophoresis device.
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4 Concluding remarks
The work shown here illustrates a microelectrophoresis
chip with an integrated sample collection scheme that is
constructed using a simple, inexpensive fabrication
methodology. Expensive micromachining equipment
associated with microtechnology is not needed. The only
materials required are glass slides, a photo-mask transparency, PIBA, and a UV source. The cost of these materials is minimal and will allow researchers to take advantage of the benefits of microscale analysis. Devices are
built in minutes and modifications to a design are made
easily by changing a photo-mask transparency. The cost
of such a mask is typically $20 US (as compared to
standard chrome masks that cost $400–$500 US).
One important point this device addresses is the issue of
post separation sample collection without the use of
expensive liquid handling instruments. The RCI presented
here provides a sample recovery method that can interface with the macro world, allowing researchers to further
process samples with traditional molecular biology techniques. For demonstration purposes, we have shown a
simple device for 1-D single lane separation of biomolecules. It should be noted that parallel lanes can be easily
integrated into this platform allowing for multiple sample
analysis. As mentioned above, various microfluidic tools
have been demonstrated using mFT. Electrophoresis and
the RCI are additional tools that can be added to the mFT
toolbox to create a versatile platform for electrophoretic
and other biological analysis methods.
© 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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