A Digital Microfluidic Approach to Homogeneous
Enzyme Assays
Elizabeth M. Miller and Aaron R. Wheeler*
Department of Chemistry, University of Toronto, 80 St. George Street, Toronto, ON, M5S 3H6, Canada, and Donnelly
Centre for Cellular and Biomolecular Research, 160 College Street., Toronto, ON, M5S 3E1, Canada
A digital microfluidic device was applied to a variety of
enzymatic analyses. The digital approach to microfluidics
manipulates samples and reagents in the form of discrete
droplets, as opposed to the streams of fluid used in
channel microfluidics. This approach is more easily
reconfigured than a channel device, and the flexibility of
these devices makes them suitable for a wide variety of
applications. Alkaline phosphatase was chosen as a model
enzyme and used to convert fluorescein diphosphate into
fluorescein. Droplets of alkaline phosphatase and fluorescein
diphosphate were merged and mixed on the
device, resulting in a 140-nL, stopped-flow reaction
chamber in which the fluorescent product was detected
by a fluorescence plate reader. Substrate quantitation was
achieved with a linear range of 2 orders of magnitude and
a detection limit of ∼7.0 × 10-20 mol. Addition of a small
amount of a nonionic surfactant to the reaction buffer was
shown to reduce the adsorption of enzyme to the device
surface and extend the lifetime of the device without
affecting the enzyme activity. Analyses of the enzyme
kinetics and the effects of inhibition with inorganic
phosphate were performed, and Km and kcat values of 1.35
µM and 120 s-1, respectively, agreed with those obtained
in a conventional 384-well plate under the same conditions
(1.85 µM and 155 s-1). A phototype device was also
developed to perform multiplexed enzyme analyses. It was
concluded that the digital microfluidic format is able to
perform detailed and reproducible assays of substrate
concentrations and enzyme activity in much smaller
reaction volumes and with higher sensitivity than conventional
methods.
Enzymatic assays are a key component in a variety of
applications, such as glucose monitoring in serum1 and screening
for phenols2 or organophosphates3 in drinking water. Relative to
other analytical techniques such as separations and mass spectrometry,
enzyme assays can be used to detect or quantify the
concentrations of small molecules in a format that is far more
compact, portable, and easy-to-use. In addition, enzyme assays
are uniquely suited for analyzing the activity of proteins (instead
of merely detecting them). A challenge for enzyme assays is in
analytical capacity and sensitivitysseveral different analyses may
be required to characterize a highly complex sample (e.g., a
cellular extract), and many replicates of each analysis must be
performed to ensure the reproducibility of the data. As the number
of analyses being performed increases, the cost of reagents and
the amount of sample required (often available in limited quantities)
become important factors. The technology of microfluidics,
which can be used to examine many samples in parallel while
consuming smaller amounts of samples and reagents than is
required for conventional-scale methods, has the potential to make
enzyme assays indispensable for a wide range of applications.4
Most microfluidic enzyme assays have been implemented via
the channel microfluidic format, in which reagents and samples
are transported through a system of enclosed micrometerdimension
channels as streams of fluid. Hadd et al.5,6 performed
some of the first enzyme assays in microchannels in a continuousflow
format and demonstrated a 4 order of magnitude reduction
in enzyme and substrate consumption over conventional methods.
However, this system relied on diffusional mixing, and reaction
times of up to 20 min were required to obtain accurate kinetic
data. With the help of a microfabricated mixer, Burke et al.7
conducted a stopped-flow enzyme assay that consumed only 6 nL
of enzyme and required less than 60 s to complete. While this
approach greatly reduces reagent consumption and analysis times
over macroscale methods, fabrication methods for such systems
are often expensive and time-consuming.8 Once a particular
arrangement of channels and reaction chambers is designed, a
great deal of time and expense is required to change the device
to suit a different application. The fabrication of multiplexed
devices becomes a daunting prospect when features such as
mixers must be incorporated, as even the simplest mixing
schemes require an integrated network of many smaller features
such as channels9 or raised ridges.10
* To whom correspondence should be addressed. E-mail: awheeler@
chem.utoronto.ca. Tel: (416) 946 3864. Fax: (416) 946 3865.
(1) Reach, G.; Wilson, G. S. Anal. Chem. 1992, 64, 381A-386A.
(2) Chen, W. J. Anal. Chim. Acta 1995, 312, 39-44.
(3) Mulchandani, A.; Chen, W.; Mulchandani, P.; Wang, J.; Rogers, K. R. Biosens.
Bioelectron. 2001, 16, 225-230.
(4) Miller, E. M.; Freire, S. L. S.; Wheeler, A. R. In Encyclopedia of Micro- and
Nanofluidics; Li, D. Q., Ed.; Springer: Heidelberg, Germany, in press.
(5) Hadd, A. G.; Raymond, D. E.; Halliwell, J. W.; Jacobson, S. C.; Ramsey, J.
M. Anal. Chem. 1997, 69, 3407-3412.
(6) Hadd, A. G.; Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1999, 71, 5206-
5212.
(7) Burke, B. J.; Regnier, F. E. Anal. Chem. 2003, 75, 1786-1791.
(8) Dittrich, P. S.; Tachikawa, K.; Manz, A. Anal. Chem. 2006, 78, 3887-3908.
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Anal. Chem. 2008, 80, 1614-1619
1614 Analytical Chemistry, Vol. 80, No. 5, March 1, 2008 10.1021/ac702269d CCC: $40.75 © 2008 American Chemical Society
Published on Web 01/26/2008
An alternative paradigm for microfluidics has recently emerged,
in which small volumes of fluid are manipulated in the form of
droplets on an open platform instead of as streams in enclosed
channels. This approach, called digital microfluidics (DMF),
exploits the electrowetting and dielectrophoresis forces generated
when an electrical potential is applied to an electrode in an
array.11-14 By applying electrical potentials to sequential electrodes,
a droplet of fluid can be dispensed from a reservoir, transported
to any position on the array, and merged with other droplets to
perform reactions.15,16 This technique has been used to actuate a
wide range of volumes (nL to µL), and unlike channel devices,
there is no sample wasted in creating a small plug for analysis. In
addition, each droplet is isolated from its surroundings rather than
being embedded in a stream of fluidsa simple method of forming
a microreactor in which there is no possibility that products will
diffuse away. Perhaps most importantly, the geometry of the array
ensures that any device design is reconfigurablessince a greater
number of paths through the device are possible, a variety of
functions can be performed without redesigning the device. There
is currently much enthusiasm for applying DMF to multiplexed
assays;17 however, it has only been applied to a few biological
applications.18-25 One reason for the lack of applications for this
technique is that fabrication of DMF devices is arduous and timeconsuming;
we recently developed a suite of rapid prototyping
methods for DMF devices,26-28 which should make the method
available to all who wish to use it.
A wide variety of channel microfluidic devices has been
developed for enzyme studies,5-7,29 but the potential of digital
devices has been largely unexplored. Taniguchi et al.18 demonstrated
the movement and merging of droplets of luciferase and
luciferin on an electrowetting-based device, but this system
required a hydrophobic liquid (oil) medium surrounding the
droplet, and no quantitative data were obtained. A two-enzyme
assay for the detection of glucose was reported by Srinivasan et
al.19, 20 that also required a silicone oil medium to achieve droplet
movement and prevent analyte adsorption to the device surface.
While this method proved to be effective for the quantitation of
glucose in a variety of biological fluids, a detailed analysis of
enzyme activity was not achieved. It would be desirable to develop
an assay that does not require oil, as it can make working with
some solvents such as ethanol or methanol impossible; it is also
likely that hydrophobic analytes may partition out of the aqueous
droplet and into the oil, resulting in incomplete reactions,
inaccurate quantitation, and cross-contamination. Jary et al.21
developed a microfluidic microprocessor for DNA repair enzyme
analysis that was just as sensitive as macroscale methods, but
with a more limited dynamic range. An ideal microfluidic assay
system would have high sensitivity, yet retain the flexibility and
dynamic range of its large-scale counterpart.
Here, we demonstrate the first application of digital microfluidic
methods to the detection and quantification of small molecules
via an enzyme assay, as well as for the study of enzyme kinetics,
with no need for liquid suspending media (e.g, silicone oil). We
show that the DMF assay has better sensitivity than macroscale
methods, but without sacrificing dynamic range. This technique
has great potential as a simple yet versatile analytical tool for
implementing enzymatic analyses on the microscale.
EXPERIMENTAL SECTION
Reagents and Materials. Alkaline phosphatase (type VII-L
from bovine intestinal mucosa), diethanolamine, magnesium
chloride, Fluorinert FC-40, sodium phosphate dibasic, and Pluronic
F-127 were purchased from Sigma Chemical (Oakville, ON,
Canada). Fluorescein diphosphate (FDP) was purchased from
Molecular Probes (Invitrogen Canada, Burlington, ON, Camada).
Parylene-C dimer was from Specialty Coating Systems (Indianapolis,
IN), and Teflon-AF was purchased from DuPont (Wilmington,
DE).
Clean room reagents and supplies included Shipley S1811
photoresist and developer from Rohm and Haas (Marlborough,
MA), solid chromium and gold from Kurt J. Lesker Canada
(Toronto, ON, Canada), CR-4 chromium etchant from Cyantek
(Fremont, CA), hexamethyldisilazane (HMDS) from Shin-Etsu
MicroSi (Phoenix, AZ), AZ-300T photoresist stripper from AZ
Electronic Materials (Somerville, NJ), and conc. sulfuric acid and
hydrogen peroxide (30%) from Fisher Scientific Canada (Ottawa,
ON, Canada). Piranha solution was prepared as a 3:1 (v/v) mixture
of sulfuric acid and hydrogen peroxide.
Device Fabrication. Digital microfluidic devices were formed
using conventional methods in the University of Toronto Emerging
Communications Technology Institute (ECTI) fabrication
facility. Glass wafers were cleaned in piranha solution (10 min)
and then coated with chromium (10 nm) and gold (100 nm) by
electron beam deposition. After rinsing and baking on a hot plate
(115 °C, 5 min), the substrates were primed by spin-coating with
HMDS (3000 rpm, 30 s) and then spin-coated with Shipley S1811
photoresist (3000 rpm, 30 s). Substrates were prebaked on a hot
plate (100 °C, 2 min) and exposed through a photomask using a
Suss Mikrotek mask aligner. Substrates were developed in MF321
(3 min) and then postbaked on a hot plate (100 °C, 1 min). After
photolithography, substrates were immersed in gold etchant
(10) Stroock, A. D.; Dertinger, S. K. W.; Adjari, A.; Mezic, I.; Stone, H. A.;
Whitesides, G. M. Science 2002, 295, 647-651.
(11) Lee, J.; Moon, H.; Fowler, J.; Schoellhammer, T.; Kim, C.-J. Sens. Actuators,
A 2002, 95, 259-268.
(12) Pollack, M. G.; Fair, R. B.; Shenderov, A. D. Appl. Phys. Lett. 2000, 77,
1725-1726.
(13) Washizu, M. IEEE Trans. Ind. Appl. 1998, 34, 732-737.
(14) Velev, O. D.; Prevo, B. G.; Bhatt, K. H. Nature 2003, 426, 515-516.
(15) Fowler, J.; Moon, H.; Kim, C.-J. Proc. IEEE Conf. Microelectromech. Syst.
2002, 97-100.
(16) Cho, S. K.; Moon, H.; Kim, C.-J. J. Microelectromech. Syst. 2003, 12, 70-
80.
(17) Mukhopadhyan, R. Anal. Chem. 2006, 78, 1401-1404.
(18) Taniguchi, T.; Torii, T.; Higuchi, T. Lab Chip 2002, 2, 19-23.
(19) Srinivasan, V.; Pamula, V. K.; Fair, R. B. Lab Chip 2004, 4, 310-315.
(20) Srinivasan, V.; Pamula, V. K.; Fair, R. B. Anal. Chim. Acta 2004, 507, 145-
150.
(21) Jary, D.; Chollat-Namy, A.; Fouillet, Y.; Boutet, J.; Chabrol, C.; Castellan,
G.; Gasparutto, D.; Peponnet, C. In NSTI Nanotech Technical Proceedings
2006, Vol. 2, pp 554-557.
(22) Chang, Y. H.; Lee, G. B.; Huang, F. C.; Chen, Y. Y.; Lin, J. L. Biomedical
Microdevices 2006, 8, 215-225.
(23) Wheeler, A. R.; Moon, H.; Kim, C.-J.; Loo, J. A.; Garrell, R. L. Anal. Chem.
2004, 76, 4833-4838.
(24) Wheeler, A. R.; Moon, H.; Bird, C. A.; Loo, R. R. O.; Kim, C.-J.; Loo, J. A.;
Garrell, R. L. Anal. Chem. 2005, 77, 534-540.
(25) Moon, H.; Wheeler, A. R.; Garrell, R. L.; Loo, J. A.; Kim, C.-J. Lab Chip
2006, 6, 1213-1219.
(26) Watson, M.; Abdelgawad, M.; Ye, G.; Yonson, N.; Trottier, J.; Wheeler, A.
R. Anal. Chem. 2006, 78, 7877-7885.
(27) Abdelgawad, M.; Wheeler, A. R. Adv. Mater. 2007, 19, 133-137.
(28) Abdelgawad, M.; Wheeler, A. R. Microfluidics and Nanofluidics; in press.
DOI: 10.1007/s10404-007-0190-3.
(29) Duffy, D. C.; Gillis, H. L.; Lin, J.; Sheppard, N. F. Jr.; Kellogg, G. J. Anal.
Chem. 1999, 71, 4669-4678.
Analytical Chemistry, Vol. 80, No. 5, March 1, 2008 1615
(60 s) followed by chromium etchant (60 s). Finally, the remaining
photoresist was stripped in AZ300T (5 min) in an ultrasonic
cleaner.
After forming electrodes, devices were coated with 2 µm of
parylene C and 50 nm of Teflon-AF. Parylene C was applied using
a vapor deposition instrument (Specialty Coating Systems), and
Teflon-AF was spin-coated (1% w/w in Fluorinert FC-40, 2000 rpm,
60 s) and then postbaked on a hot plate (160 °C, 10 min). To
actuate droplets, the polymer coatings were locally removed from
the contact pads by gentle scraping with wafer tweezers. In
addition to patterned devices, unpatterned indium-tin oxide (ITO)coated
glass substrates (Delta Technologies Ltd, Stillwater, MN)
were coated with Teflon-AF (50 nm, as above).
Droplet Actuation. Each device was assembled with an
unpatterned ITO/glass top plate and a patterned bottom plate
separated by a spacer formed from one piece of double-sided tape
(∼70 µm thick). The 140-nL droplets in devices with this
arrangement had a diameter of ∼1.6 mm (∼40% of the size of a
well in a 384-well plate). As described previously,11,16,24 droplets
were sandwiched between the two plates and actuated by applying
electric potentials between the top electrode and sequential
electrodes on the bottom plate. Applied potentials (60-80 VRMS)
were generated by amplifying the output of a function generator
operating at 18 kHz. Driving potentials were applied manually to
exposed contact pads on the bottom plate surface. Droplet
actuation was monitored and recorded by a CCD camera mated
to a stereomicroscope with fluorescence imaging capability
(Olympus Canada, Markham, ON, Canada). All devices had 1 mm
× 1 mm actuation electrodes, with an interelectrode gap of 40-
75 µm.
Analysis of Enzyme Activities and Kinetics. Alkaline phosphatase
solutions were prepared in 10 mM diethanolamine (DEA)
buffer, pH 10.1, containing 1 mM MgCl2 and 0.1% (w/v) Pluronic
F-127. Enzyme concentrations were 6 U/mL for standard curve
experiments and for observation through the microscope and 0.21
U/mL for kinetic analyses. Fluorescein diphosphate solutions of
various concentrations were prepared in 10 mM DEA buffer. For
enzyme inhibition experiments, fluorescein diphosphate solutions
containing various concentrations of inorganic phosphate were
also prepared. For all quantitative experiments, a digital microfluidic
device was manually positioned on the top of a 384-well
microtiter plate, and a 650-nL reservoir droplet of enzyme or
substrate solution was pipetted onto each of two large electrodes
(2 mm × 2 mm). As depicted in Figure 1, 70-nL droplets of each
reagent were actively dispensed from the reservoirs (as described
previously16). Briefly, two electrodes adjacent to the reservoir were
actuated sequentially to form a finger of fluid extending onto the
array; upon simultaneous actuation of the target electrode and
the reservoir electrode; the finger was split, resulting in a
dispensed droplet with a volume defined approximately by the
electrode and spacer dimensions (1 mm × 1 mm × 70 µm). The
two dispensed reagent droplets were then merged and mixed by
moving the coalesced droplet around a loop of six actuation
electrodes for 15 s. After mixing, the device (still positioned on
the microtiter plate) was inserted into a PheraStar multiwell plate
reader (BMG Labtech, Durham, NC) for fluorescence detection
(λex ) 485 nm; λem ) 520 nm; focal height, 15.0 mm; gain, 1462
or 90 for substrate quantitation and kinetics experiments, respectively).
After each assay, devices were rinsed in DEA buffer and
DI water and allowed to dry. Because of the finite time required
Figure 1. Fluorescent enzymatic assay on a DMF device. A droplet containing FDP was dispensed from the reservoir on the right (a) and a
droplet of AP was dispensed from the reservoir on the left (a, b). When the droplets were merged under fluorescent illumination, the product
was observed at the interface of the droplets (c). After active mixing, the reaction proceeded to completion (d). Lines have been added to the
images in (a, b) to indicate the location and movement of droplets.
1616 Analytical Chemistry, Vol. 80, No. 5, March 1, 2008
for mixing of the reagents, insertion of devices into the plate
reader, and measurement of the fluorescent signals, kinetic assays
were conducted at low concentrations of enzyme and substrate,
making the linear approximation of the initial rate more accurate.
The concentration detection limits were determined by sequentially
diluting the substrate solution until the measured signal
intensity was equal to the background plus 3× the standard
deviation of the background measured from 10 droplets not
containing the fluorophore. For all experiments, 3-5 trials were
conducted to evaluate reproducibility of the method.
For comparison, all assays implemented by digital microfluidics
(in 140-nL droplets, as described above) were also carried out in
384-well microtiter plates. For these conventional-scale assays, 10
µL each of the enzyme and substrate solutions were pipetted into
wells and the plate was shaken by the plate reader to ensure
mixing of the reagents before detection. All experiments in
microtiter plates used the same enzyme and substrate solutions
used on digital microfluidic devices to ensure that each assay had
similar dilution and mixing conditions, and 3-5 replicates of each
measurement were performed for error analysis.
RESULTS AND DISCUSSION
Device Optimization. A common difficulty in the application
of DMF devices to proteomic assays lies in achieving consistent
actuation of droplets of protein solutions. Droplet actuation by
means of electrowetting requires a hydrophobic electrode surface,
and protein deposition on this hydrophobic surface reduces the
efficiency of actuation and the lifetime of the device.30 The alkaline
phosphatase (AP) used here had sufficient activity such that
substrate turnover was observed when the enzyme was present
at concentrations of 6 U/mL (2 µg/mL) or lesssa concentration
range for proteins that has been reported to be compatible with
DMF.20,30
In preliminary work, we noticed that, even when using low
concentrations of alkaline phosphatase, some enzyme adsorbed
to the Teflon surface during the course of experiments, which
caused the devices to fail (i.e., failed devices could no longer
support droplet movement) after a small number of assays. In an
attempt to extend the device lifetimes and reduce crosscontamination,
we evaluated the effect of supplementing the
carrier buffers with Pluronic F-127, a triblock copolymer and
nonionic surfactant that has been shown to reduce nonspecific
adsorption of proteins to hydrophobic surfaces.31-33 In initial trials,
we observed that droplets containing low concentrations of
Pluronic F-127 were reliably moveable by DMF, although they
moved less rapidly than droplets not containing the surfactant.
When used in enzyme assays, the strategy was remarkably
successful at extending device lifetimessin one experiment using
alkaline phosphatase at 6 U/mL (2 µg/mL) with 0.1% w/v Pluronic
F-127, a single device was used for 21 consecutive assays, with
no noticeable degradation of device performance. Between assays,
a simple rinse with buffer was sufficient to maintain droplet
actuation, and no contribution to the error in subsequent trials
was observed (i.e., no evidence of cross-contamination). In
ongoing work, we are evaluating the optimal conditions (concentration
and type of Pluronic, etc.) for reducing unwanted adsorption
in DMF;34 we used 0.1% w/v F-127 for all of the results
reported here.
Enzyme Activity. In order to determine the potential of the
new DMF method for substrate quantitation and other enzymatic
analyses, a standard curve was generated and compared to a well(30)
Yoon, J.-Y.; Garrell, R. L. Anal. Chem. 2003, 75, 5097-5102.
(31) Amiji, M.; Park, K. Biomaterials 1992, 13, 682-692.
(32) Ng, C. L.; Lee, H. K.; Li, S. F. Y. J. Chromatogr. 1994, 659, 427-434.
(33) McClain, M. A.; Culbertson, C. T.; Jacobson, S. C.; Allbritton, N. L.; Sims,
C. E.; Ramsey, J. M. Anal. Chem. 2003, 75, 5646-5655.
(34) Luk, V.; Mo, G. C. H.; Wheeler, A. R. Sumitted to Langmuir.
Figure 2. FDP calibration curves. On a 384-well plate (a) and a
DMF device (b), the fluorescence response was linear over a range
of 2 orders of magnitude of substrate concentration. Error bars are
(1 SD. In both experiments, the error in the data points ranged from
<2% to ∼10% RSD.
Figure 3. Effect of Pluronic F-127 on enzyme assays. An APFDP
reaction was allowed to proceed to completion in a well plate
under three sets of conditions: (1) without any Pluronic added to the
reaction mixture, (2) with Pluronic added to the wells after the reaction
was completed, and (3) with Pluronic added to the wells at the start
of the reaction. Error bars are (1 SD. While the presence of the
polymer does reduce the fluorescent signal at the detector, the
reduction is the same whether the Pluronic is added at the beginning
of the reaction or at completion. The activity of the enzyme appears
to be unaffected.
Analytical Chemistry, Vol. 80, No. 5, March 1, 2008 1617
plate experiment under the same conditions (Figure 2). Both
methods had a dynamic range of ∼2 orders of magnitude, which
appears to be limited only by the detectorsthere is no marked
difference between the data generated by the two methods. The
concentration detection limits of the DMF and well-plate methods
were determined to be approximately 1.0 and 0.50 pM FDP,
respectively. These concentration data represent mass detection
limits of 7.0 × 10-20 and 5.0 × 10-18 mol, respectivelysan
improvement of nearly 2 orders of magnitude in the microfluidic
system. This increase in sensitivity is likely a function of the
placement of the device within the plate readersthe DMF device
is closer to the detector than the fluid at the bottom of the well.
In addition, the gold surface of the electrodes on DMF devices
reflects the emitted light rather than absorbing it (as in the black
wells), so more fluorescence is likely collected from the DMF
device. The dispensing of 70-nL droplets on the microscale device
appears to be nearly as reproducible as dispensing 10-µL aliquots
via micropipet on the macroscale; the error in each data point is
within ∼10% RSD in both experiments.
The effect of the Pluronic additive on the assay was also
examined. An AP-FDP reaction was carried out in a 384-well plate
under three sets of conditions: (1) without any Pluronic added
to the reaction mixture, (2) with Pluronic added to the wells after
the reaction was completed, and (3) with Pluronic added to the
wells at the start of the reaction. As shown in Figure 3 (comparing
conditions 1 and 3), the presence of the additive decreases the
fluorescent signal measured by the detector by ∼30%. As shown
by comparing conditions 2 and 3, the decrease in the fluorescent
signal is identical for cases when Pluronic is added at the
beginning or end of the reaction, which indicates that this effect
is not caused by a decrease in the amount of product. Therefore,
the effect is likely caused by interaction of the fluorescent product
with Pluronic molecules rather than by a reduction in enzyme
activity caused by the surfactant. Regardless, the effect is small
and does not cause problems for conducting enzyme assays.
Enzyme Kinetics. After demonstrating the compatibility of
DMF with fluorogenic enzyme assays, we applied the technique
to measuring Michaelis-Menten enzyme kinetics. Since ∼30 s
elapses from the time the reagent droplets are merged to the time
that the first fluorescence data points can be collected, experimental
conditions were optimized such that the rate at 30 s was
similar to the initial rate. As shown in Figure 4a, it was determined
that an AP concentration of 0.21 U/mL met these conditions. A
series of experiments was then performed, implemented on DMF
devices and in 384-well plates, in which the enzyme concentration
was held constant at 0.21 U/mL while the substrate concentration
was varied and initial velocities were calculated. Five replicates
were run at each substrate concentration to determine the
precision of each method. The initial velocities were then plotted
against substrate concentrations in a double-reciprocal Lineweaver-Burk
plot (Figure 4b).
For both formats (macro- and microscale), the LineweaverBurk
plot was linear over a range of ∼1 order of magnitude. The
DMF data appear slightly more reproducible than the 384-well
plate (errors in the data from the DMF device ranged from 2 to
10% RSD, while errors in the data from the macroscale method
varied from 7 to 17% RSD). The values of Km, Vmax, and kcat
calculated from these plots are shown in Table 1. In both cases,
the calculated constants are similar to those typically reported
Figure 4. Study of enzyme kinetics on a DMF device. After droplets
containing enzyme and substrate (2.5 nM) were merged and mixed,
the fluorescent product was monitored over time (a). The initial rate
was measured at several substrate concentrations and reported as
a Lineweaver-Burk plot (filled circles) (b) and compared to a
macroscale experiment in a 384-well plate (open circles). Error bars
are (1 SD.
Table 1. Michaelis-Menten Kinetic Variables for
Alkaline Phosphatase
384-well plate DMF device
Vmax 54.8 ( 5.7 nM/s 42.5 ( 7.8 nM/s
Km 1.85 ( 0.2 µM 1.35 ( 0.3 µM
kcat 155 ( 16 s-1 120. ( 20 s-1
Figure 5. Inhibition of AP with Pi. The concentration of FDP was
held constant at either 10 or 5 µM and the concentration of Pi varied
from 0 to 45 µM. The rate of reaction is reduced with each increase
in inhibitor concentration, as the inorganic phosphate competes with
the substrate for the active sites on the enzyme.
1618 Analytical Chemistry, Vol. 80, No. 5, March 1, 2008
for alkaline phosphatase (kcat ∼100).35 The DMF-determined value
of kcat is slightly lower than that obtained by using well platess
this is likely due to the longer time lag between mixing and
measurement. However, there is excellent overall agreement
between the values calculated from the micro- and macroscale
systems, which suggests that there is great potential for using
DMF for the analysis of enzyme kinetics.
In addition to Michaelis-Menten analyses, we evaluated the
suitability of DMF for implementing enzyme inhibition studies.
In these experiments, inorganic phosphate (Pi) at various concentrations
was added to the droplet containing FDP. Pi is a wellknown
competitive inhibitor of alkaline phosphatase,36 and as
shown in Figure 5, the rate of reaction is slowed with each increase
in Pi concentration. These qualitative results are particularly
exciting, as in the future, we plan to use DMF to carry out
quantitative enzyme inhibitor screens to identify new reagents with
pharmaceutical activities. As described above, the advantages for
digital microfluidics of reduced reagent consumption and increased
assay sensitivity make this plan a promising one.
Multiplexed Devices. Recently, we designed and fabricated
a DMF device for multiplexed analysis, shown in Figure 6. This
device contains six identical reaction platforms connected in series,
which facilitates simultaneous control of six separate reactions
using a single program of driving potentials. The spacing between
platforms is 9 mm, the same as the interwell pitch on a 96-well
plate, allowing all six devices to be simultaneously monitored
within the plate reader. We are currently optimizing the operating
parameters for this device; when completed, we expect the new
design (or a similar one) will be characterized by improved speed
and precision of this method. Each measurement will have a builtin
redundancy, eliminating the need to run replicates and reducing
the amount of time required to perform an analysis. Devices such
as the one pictured in Figure 6 are moving us one step closer to
realizing a high-throughput microfluidic platform for multiplexed
screening.
CONCLUSION
Enzymatic assays are widely used in carrying out chemical
analyses on the microscale. We have demonstrated that the digital
microfluidic paradigm is well-suited to these assays, without the
flow control, mixing, and fabrication problems encountered in
channel microfluidic devices. In terms of small-molecule detection
and quantitation, the digital microfluidic device performed comparably
to the macroscale method, with standard errors of 10%
or less. An increase in sensitivity of nearly 2 orders of magnitude
was achieved, while decreasing sample and reagent consumption
by 2.5 orders of magnitude. Reproducible and accurate kinetic
assays were implemented via DMF, yielding kinetic constants that
agree with those reported in the literature. Protein adsorption to
device surfaces can be controlled with the addition of a Pluronic
polymer to the reaction buffersa silicone oil medium is not
required. Finally, we conclude that screening of inhibitors and
multiplexed analyses are possible, which will decrease analysis
times and enable us to achieve a truly high-throughput enzymatic
assay on the microscale.
ACKNOWLEDGMENT
We acknowledge the Natural Sciences and Engineering
Council (NSERC) and the Canada Foundation for Innovation (CFI)
for financial support. A.R.W. thanks the CRC for a Canada
Research Chair.
Received for review November 2, 2007. Accepted
December 9, 2007.
AC702269D
(35) Craig, D. B.; Arriaga, E. A.; Wong, J. C. Y.; Lu, H.; Dovichi, N. J. J. Am.
Chem. Soc. 1996, 118, 5245-5353.
(36) McComb, R. B.; Bowers Jr. G. N.; Posen, S. Alkaline Phosphatase; Plenum
Press: New York, 1979.
Figure 6. Multiplexed DMF device. Picture of a device designed for simultaneous control of six independent DMF-driven assays (a). A droplet
of fluorescein is dispensed from a reservoir on several platforms via a single set of electrical contacts (b).
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