Measurement of enzyme kinetics using a continuous-flow microfluidic system

Article abstract of DOI:10.1021/ac034155b

This paper describes a microanalytical method for determining enzyme kinetics using a continuous-flow microfluidic system. The analysis is carried out by immobilizing the enzyme on microbeads, packing the microbeads into a chip-based microreactor (volume ～1.0 nL), and flowing the substrate over the packed bed. Data were analyzed using the Lilly-Hornby equation and compared to values obtained from conventional measurements based on the Michaelis-Menten equation. The two different enzyme-catalyzed reactions studied were chosen so that the substrate would be nonfluorescent and the product fluorescent. The first reaction involved the horseradish peroxidase-catalyzed reaction between hydrogen peroxide and N-acetyl-3,7-dihydroxyphenoxazine (amplex red) to yield fluorescent resorufin, and the second the β-galactosidase-catalyzed reaction of nonfluorescent resorufin-β-D-galactopyranoside to yield D-galactose and fluorescent resorufin. In both cases. the microfluidics-based method yielded the same result obtained from the standard Michaelis-Menten treatment. The continuous-flow method required ～10μL of substrate solution and 109 enzyme molecules. This approach provides a new means for rapid determination of enzyme kinetics in microfluidic systems, which may be useful for clinical diagnostics, and drug discovery and screening.

Full text of DOI:10.1021/ac034155b

Anal. Chem. 2003, 75, 3161-3167
Measurement of Enzyme Kinetics Using a
Continuous-Flow Microfluidic System
Gi Hun Seong,† Jinseok Heo, and Richard M. Crooks*
Department of Chemistry, Texas A&M University, P.O. Box 30012, College Station, Texas 77842-3012
croanalytical systems awaits development of methods that capital-
This paper describes a microanalytical method for deter-
mining enzyme kinetics using a continuous-flow microf-
luidic system. The analysis is carried out by immobilizing
the enzyme on microbeads, packing the microbeads into
a chip-based microreactor (volume ∼1 .0 nL), and flowing
the substrate over the packed bed. Data were analyzed
using the Lilly-Hornby equation and compared to values
obtained from conventional measurements based on the
Michaelis-Menten equation. The two different enzyme-
catalyzed reactions studied were chosen so that the
substrate would be nonfluorescent and the product fluo-
rescent. The first reaction involved the horseradish per-
oxidase-catalyzed reaction between hydrogen peroxide
and N-acetyl-3 ,7 -dihydroxyphenoxazine (amplex red) to
yield fluorescent resorufin, and the second the â-galac-
tosidase-catalyzed reaction of nonfluorescent resorufin-
ize upon these enhancements without introducing complex
problems that outweigh the advantages. Here, we address this
need by describing an analytical method for measuring enzyme
kinetics using a continuous-flow microfluidic system. In this
approach, a microfluidic channel is filled with enzyme-modified
microbeads, and then a nonfluorescent substrate is moved over
the bead bed by convective flow. It is possible to correlate the
enzyme reaction rate to the appearance of the fluorescent product
of the enzyme-catalyzed reaction. Accordingly, this method is
potentially useful for high-throughput screening of enzymatic
activity.
The first report of using a microfluidic system for analyzing
the kinetics of an enzymatic reaction was provided by Hadd et
al.,12 who used computer-controlled electrokinetic transport for
diluting and mixing reagents. Specifically, precise concentrations
of a fluorogenic substrate were mixed with â-galactosidase, and
the kinetics of the reaction was obtained by monitoring the
fluorescence of the hydrolysis product. More recently, Duffy et
al.13 developed a centrifugal microfluidic system capable of
carrying out multiple simultaneous homogeneous enzymatic
assays using colorimetric detection.
Most previous quantitative reports of enzyme activity measured
using microfluidic systems have been carried out in homogeneous
solutions.14-18 However, heterogeneous assays using immobilized
enzymes have the potential for providing the following advan-
tages: ease of enzyme placement within arrays, ability to assay
membrane proteins (which of course is not possible in homoge-
neous solution), simplification of enzyme recycling, and continu-
ous-flow analysis. As discussed later, a potential drawback of our
approach is that immobilization of enzymes may alter their
â-D-galactopyranoside to yield D-galactose and fluorescent
resorufin. In both cases. the microfluidics-based method
yielded the same result obtained from the standard
Michaelis-Menten treatment. The continuous-flow method
required ∼1 0 µL of substrate solution and 1 0 9 enzyme
molecules. This approach provides a new means for rapid
determination of enzyme kinetics in microfluidic systems,
which may be useful for clinical diagnostics, and drug
discovery and screening.
Microfluidic chip-based devices have been developed for a
number of important assays, including DNA analysis,1,2 protein
analysis,3,4 and immunoassays.5-7 During the past decade it was
demonstrated that, compared to traditional analytical instruments,
the microfluidic platform provides a means for reducing both
analysis time and the amount of reagent necessary to perform
each analysis.8-11 However, profitable commercialization of mi-
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(10) Harrison, D. J., van den Berg, A., Eds. Micro Total Analysis Systems ′98;
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(12) Hadd, A. G.; Raymond, D. E.; Halliwell, J. W.; Jacobson, S. C.; Ramsey, J.
M. Anal. Chem. 1 9 9 7 , 69, 3407-3412.
* Corresponding author: (voice) 979-845-5629; (fax) 979-845-1399; (e-mail)
crooks@tamu.edu.
† Present address: Department of Chemistry and Applied Chemistry, Hanyang
University, Ansan, Kyunggi-do 426-791, Korea.
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10.1021/ac034155b CCC: $25.00 © 2003 American Chemical Society
Published on Web 05/14/2003
Analytical Chemistry, Vol. 75, No. 13, July 1, 2003 3161

intrinsic kinetic rates. Interestingly, only a few reports describe
the use of biomolecules immobilized either directly on the walls
of microchannels or on a support contained within the channel.
For example, a very early report from Karube and co-workers
described a sensor for glucose detection based on immobilization
of glucose oxidase on the walls of a micromachined capillary.19
More recently, Harrison et al. showed that a microfluidic device
containing trypsin immobilized on beads could be used to digest
proteins prior to analysis by mass spectrometry.20 Mao et al.
reported a method for immobilizing enzymes on fluid bilayers
supported on the walls of microfluidic channels and then evaluat-
ing their kinetics in one shot using laminar flow-controlled
dilution.21 Finally, Peterson et al. reported the preparation of
reactive porous monoliths within microchannels, their use as
supports for the immobilization of trypsin, and the characterization
of the resulting conjugates for the digestion of proteins.22
In previous reports, we described microbead-based microfluidic
systems for carrying out multiple, sequential enzymatic reactions
and DNA hybridization analyses.23,24 A key finding of these studies
is that microbeads offer the convenience of fluidic handling, a high
degree of synthetic and characterization flexibility, very high
surface-to-volume ratio compared to open-channel microfluidic
devices, and the capacity to efficiently mix reagents even under
laminar flow conditions.
The intrinsic kinetics of soluble enzymes in homogeneous
solution are generally described by the Michaelis-Menten
relationship.25 However, the kinetics for immobilized enzymes may
be different from those measured for the same enzyme in solution
because of diffusional restrictions or interactions with the support.
The kinetics of immobilized biocatalysts are often referred to as
the apparent kinetics. Under steady-state conditions, the apparent
kinetic parameters in packed-bed enzyme reactors are usually
calculated using a formalism known as the Lilly-Hornby model.26
In the experiments reported here, horseradish peroxidase
(HRP) was immobilized on microbeads via streptavidin-biotin
conjugation, and the kinetics of the HRP-catalyzed reaction
between hydrogen peroxide and N-acetyl-3,7-dihydroxyphenox-
azine (amplex red) was monitored under continuous-flow condi-
tions. The results compare favorably to literature values and to
values we obtained using conventional analysis methods. To
investigate the generality of this approach, the â-galactosidase (â-
Streptavidin-Coated Microspheres (Bangs Laboratories, Fishers,
IN) that were found by optical microscopy to be 15.5 µm in
diameter. Biotin-labeled horseradish peroxidase, biotin-labeled
â-galactosidase, and sodium resorufin (7-hydroxy-3H-phenoxazin-
3-one) were purchased from the Sigma Chemical Co. (St. Louis,
MO). Amplex red (N-acetyl-3,7-dihydroxyphenoxazine) and re-
sorufin â-D-galactopyranoside were obtained from Molecular
Probes (Eugene, OR). Hydrogen peroxide (30%) and dimethyl
sulfoxide (DMSO) were obtained from EM Science (Gibbstown,
NJ). All chemicals were of reagent grade quality or better. Milli-Q
(Millipore) water (18 MΩ‚cm) was used to prepare aqueous
solutions.
Fabrication of Microfluidic Devices. The fabrication of
microfluidic devices and the associated weir structures were
described in our previous reports.23,24 Briefly, positive photoresist
was spin-coated onto a glass slide and illuminated with UV
radiation through a mask for 5 min to create a master. A slit-type
photomask having a width of 100 µm was aligned on the glass
master to fabricate weirs within the microchannels. After UV
exposure for 5 min, the glass master was developed in 60% AZ421K
solution for 30 s. PDMS polymer molds were prepared by casting
a PDMS prepolymer against a positive photoresist master and
curing at 65 °C for 2 h. The PDMS mold was irreversibly sealed
to a clean glass substrate after both were treated with an oxygen
plasma (60 W, model PDC-32G, Harrick Scientific, Ossining, NY)
for 60 s. The depth and width of the microchannel and weir were
determined using a Veeco Dektak 3 profilometer (Veeco Instru-
ments, Plainview, NY).
Enzyme Immobilization on Microbeads. HRP and â-Gal
were conjugated to biotin via amide coupling of an aminocaproyl
spacer. HRP-biotin (240 units/ mg) and â-Gal-biotin (350-700
units/ mg) contain 2-3 mol of biotin/ mol of HRP and 2-4 mol of
biotin/ mol of â-Gal, respectively. The conjugation of streptavidin-
coated microbeads with the biotinylated enzymes was carried out
using the following procedure. A 30-µL sample of stock beads (4.8
× 106 bead/ mL) were rinsed in 200 µL of phosphate buffered
saline (PBS) solution (pH 7.4, 150 mM NaCl, 4 mM KCl, 8.1 mM
Na2HPO4, and 1.47 mM KH2PO4) and centrifuged at 4000 rpm
for 5 min. The microbead pellet was resuspended in 30 µL of PBS,
and then 30 µL of the biotinylated enzymes (1 mg/ mL) in PBS
(pH 7.4) was added to the microbeads to yield the microbead-
enzyme complex. The mixture of microbeads and biotinylated
enzymes was incubated for 2 h at room temperature (18-25 °C)
with gentle mixing. After conjugation, the mixture was centrifuged
to remove unreacted biotinylated enzymes and resuspended in
500 µL PBS. These microbead-enzyme complexes were stored
at 4 °C prior to use.
Gal) catalyzed hydrolysis of resorufin â-D-galactopyronoside (RBG)
was also studied.
EXPERIMENTAL SECTION
Chemicals. Positive photoresist (AZP4620) and developer
solution (AZ421K) were obtained from the Clariant Co. (Somer-
ville, NJ). Poly(dimethylsiloxane) (PDMS) microfluidic devices
were fabricated by curing the prepolymer components of Sylgard
184 (Dow Corning, Midland, MI). Microbeads were Proactive
Enzyme Kinetic Assays. To carry out enzyme assays in
continuous-flow microfluidic systems, microbead-enzyme com-
plexes were packed into on-chip microreactors using pump-driven
pressure. Stock solutions of amplex red and RBG were prepared
in DMSO and stored at -20 °C. Immediately before use, the
thawed stock solutions were diluted in buffer solution. Various
concentrations of H2O2 (in 50 mM Tris-HCl plus 10 µM amplex
red, at pH 7.4) and RBG (in 100 mM Tris-HCl, 2 mM KCl, and
0.1 mM MgCl2, at pH 7.8) were introduced into the microchannels
at various flow rates using a microsyringe pump. Optical and
fluorescence images of the enzyme reactions were acquired with
(19) Murakami, Y.; Takeuchi, T.; Yokoyama, K.; Tamiya, E.; Karube, I.; Suda,
M. Anal. Chem. 1 9 9 3 , 65, 2731-2735.
(20) Wang, C.; Oleschuk, R.; Ouchen, F.; Li, J.; Thibault, P.; Harrison, D. J. Rapid
Commun. Mass Spectrom. 2 0 0 0 , 14, 1377-1383.
(21) Mao, H.; Yang, T.; Cremer, P. S. Anal. Chem. 2 0 0 2 , 74, 379-385.
(22) Peterson, D. S.; Rohr, T.; Svec, F.; Frechet, J. M. J. Anal. Chem. 2 0 0 2 , 74,
4081-4088.
(23) Seong, G. H.; Zhan, W.; Crooks, R. M. Anal. Chem. 2 0 0 2 , 74, 3372-3377.
(24) Seong, G. H.; Crooks, R. M. J. Am. Chem. Soc. 2 0 0 2 , 124, 13360-1361.
(25) Seigel, I. H. Biochemical Calculations; Wiley: New York, 1976.
(26) Lilly, M. D.; Hornby, W. E.; Crook, E. M. Biochem. J. 1 9 6 6 , 100, 718-723.
3162 Analytical Chemistry, Vol. 75, No. 13, July 1, 2003

Figure 1. (A) Schematic illustration of the microfluidic device used
for analysis of enzyme kinetics. (B) Optical image of a microchamber
after packing with microbeads. The regions labeled M and W are,
respectively, the microreactor and weir.
a fluorescence microscope (Nikon Eclipse TE 300, Nikon Co.,
Tokyo, Japan), equipped with band-pass filters (a cube-type XF
102-2 filter set, Omega Optical, Inc., Brattleboro, VT), a 100-W
mercury lamp, and a charge-coupled device camera (Photometrics
Ltd., Tucson, AZ). Fluorescence images were analyzed using V++
precision digital imaging software (Digital Optics, Auckland, New
Zealand).
To compare results of continuous-flow, on-chip enzyme assays
with a conventional homogeneous assay method, the HRP-
catalyzed reaction between H2O2 and amplex red, and the
hydrolysis of â-Gal and RBG, were performed by monitoring the
fluorescence of resorufin at 563 nm using a conventional spec-
trofluorometer. The fluorescence signal was calibrated using a
standard solution of resorufin.
Figure 2. (A) HRP-catalyzed reaction between nonfluorescent
amplex red and H2O2 to yield fluorescent resorufin. (B) Fluorescence
micrograph of the microreactor during continuous-flow operation. The
substrate solution contained 5 µM H2O2 and 10 µM amplex red in 50
mM Tris-HCl buffer (pH 7.4) and was introduced into the microreactor
from left to right. Flow rate, 0.5 µL/min. (C) Normalized fluorescence
intensity line scans obtained at the locations indicated by the dashed
lines in (B). The excitation and maximum emission wavelengths were
563 and 587 nm, respectively.
RESULTS AND DISCUSSION
A schematic illustration of the microfluidic device used in this
study is shown in Figure 1A. The device was fabricated from
PDMS using standard photolithographic and replica molding
methods27 (fabrication details are provided in the Experimental
Section). The key components of the device are the microreactors
(labeled M in Figure 1A) and weirs (labeled W in Figure 1A).
The width and depth of the weirs ranged from 40 to 60 µm and
7 to 12 µm, respectively. Therefore, microspheres having a
diameter of 15.5 µm are retained within microreactors (Figure
1B). Microspheres were coated with streptavidin, a tetrameric
protein having four biotin binding sites, so that nucleic acids or
proteins modified with biotin could be conjugated to the micro-
sphere surface. Microspheres were introduced into microreactor
using pump-driven pressure. The volume of the microreactor was
1.0 nL, and 250-300 microbeads could be packed into each
microreactor.
nonspecific adsorption of proteins with 1 mg/ mL BSA in PBS
solution for 3 h at room temperature. Biotin-labeled HRP was then
conjugated to the streptavidin-coated microspheres, and these
modified beads were packed into the microreactor. Aqueous
solutions containing 5 µM H2O2 and 10 µM amplex red in 50 mM
Tris-HCl buffer (pH 7.4) were introduced into the main channel
at a flow rate of 0.5 µL/ min from the inlet reservoir. Figure 2B
shows a fluorescence image of the resulting HRP-catalyzed
reaction in the continuous-flow microreactor. Fluorescence line
scans (Figure 2C) taken across the inlet and outlet streams
(obtained at positions a and b, respectively, in Figure 2B) indicate
that amplex red is a fluorogenic substrate having very low
background fluorescence but that its fluorescent product, resoru-
fin, exhibits strong fluorescence.
Enzyme kinetics were evaluated using the Lilly-Hornby
model, which was developed for packed-bed reactor systems.26
This model is embodied in eq 1. Here, f is the fraction of substrate
Oxidase enzymes, and particularly HRP, are used in a wide
variety of bioassays. Accordingly, we chose to benchmark this
continuous-flow study of enzyme kinetics using the HRP-catalyzed
reaction between H2O2 and amplex red. In the presence of HRP,
amplex red reacts with H2O2 in a 1:1 stoichiometry to produce
highly fluorescent resorufin (Figure 2A).28
f[A0] ) C/ Q + Km(app) ln(1 - f)
(1)
converted to product during the reaction, Q is the flow rate of
the substrate, [A0] is the initial substrate concentration, C is the
reaction capacity of the microreactor, and Km(app) is the apparent
Michaelis constant. The value of f was determined experimentally
by subtracting the fluorescence signal in the inlet stream from
the fluorescence signal in the outlet stream of the microreactor.
Before carrying out a continuous-flow study of enzyme kinetics,
the inner walls of the microdevice were passivated against
(27) Duffy, D. C.; McDonald, J. C.; Schueller, O. J. A.; Whitesides, G. M. Anal.
Chem. 1 9 9 8 , 70, 4974-4984.
(28) Zhou, M.; Diwu, Z.; Panchuk-Voloshina, N.; Haugland, R. P. Anal. Biochem.
1 9 9 7 , 253, 162-168.
Analytical Chemistry, Vol. 75, No. 13, July 1, 2003 3163

Figure 4. Analysis of data obtained under continuous-flow condi-
tions for the HRP-catalyzed reaction between H2O2 and amplex red
to yield fluorescent resorufin. (A) Plot of 1/C (C is the microreactor
capacity) vs 1/Q (Q is the flow rate). (B) Plot of Km(app), the apparent
Michaelis constant, vs Q.
Table 1. Effect of Flow Rate, Q, on KM(app) and the
Reactor Capacity, C, of the Microreactor for the
HRP-Catalyzed Reaction
Q
Km(app)
(µM)
C
(µL/ min)
(fmol/ min)
0.2
0.5
1.0
1.5
2.32
6.25
10.3
13.0
36.4
93.5
184
245
Figure 3. Analysis of data obtained under continuous-flow condi-
tions for the HRP-catalyzed reaction between H2O2 and amplex red
to yield fluorescent resorufin. The HRP-functionalized microbeads
were packed in a microreactor like that shown in Figure 2B. (A) Plots
of the concentration of H2O2 converted to product vs the initial H2O2
concentration as a function of flow rate (Q). The flow rates ranged
from 0.20 to 1.5 µL/min. (B) Plots of the concentration of H2O2
converted to product vs flow rate as a function of the concentrations
of H2O2. The concentrations of H2O2 ranged from 1.0 to 10.0 µM.
(C) Plots of f[A0] vs -ln(1 - f). The flow rates ranged from 0.20 to
1.5 µL/min.
cence intensity. In this study, the initial H2O2 concentrations
ranged from 1 to 10 µM and the amplex red concentration was
held constant at 10 µM. These solutions were flowed through the
microreactor at rates ranging from 0.2 to 1.5 µL/ min.
Panels A and B of Figure 3 show substrate concentration and
flow rate effects, respectively, on the HRP-catalyzed hydrolysis
of H2O2. From these data, linear plots of f[A0] versus -ln(1 - f)
were obtained (Figure 3C). Values for Km(app) and the reaction
capacity of the microreactor (C), as a function of Q, were derived
from a weighted least-squares fit to the points shown in Figure
3C. The results are summarized in Table 1. The maximum reaction
capacity, Cmax, of the microreactor can then be obtained from these
data.29 Specifically, a plot of 1/ C versus 1/ Q (Figure 4A) yields a
Because the appearance of resorufin and the disappearance of
H2O2 are related by a 1:1 stoichiometry (Figure 2A), the turnover
of H2O2 could be evaluated from calibration curves obtained from
known concentrations of resorufin and its corresponding fluores-
3164 Analytical Chemistry, Vol. 75, No. 13, July 1, 2003

Cmax value of 9.03 pmol/ min. The value of Km(app) should approach
the value of the true Michaelis constant of the free enzyme, Km,
at zero flow rate if only mass-transfer effects are responsible for
masking the intrinsic enzyme kinetics. This assumption was tested
by plotting the Km(app) values obtained against Q (Figure 4B). The
intercept on the vertical axis yields a zero-flow Km(app) value
(
K 0m(app)) of 1.51 µM. As discussed next, this value compares
favorably with measurements of Km determined using standard
methods.
Enzyme kinetics in homogeneous solution were evaluated
using the Michaelis-Menten equation (eq 2). This equation
V0 ) Vmax[S]/ (Km + [S])
(2)
relates the initial rate of the enzyme reaction, V , to the concentra-
0
tion of substrate, [S], a maximum rate, Vmax, corresponding to
the velocity of the reaction when the active sites of the enzyme
are saturated with substrate, and the Michaelis constant, Km. The
values of Km for HRP were determined using the following
procedure. First, the background fluorescence was measured 100
s after mixing various concentrations of H2O2 with 10 µM amplex
red. Second, 0.01 unit/ mL HRP in 50 mM Tris-HCl (pH 7.4) was
added to these solutions, and then fluorescence intensity was
measured for 500 s. V was determined for each H2O2 concentra-
0
tion (0.1-5.0 µM) from the initial slopes of plots of normalized
fluorescence intensity as a function of time (Figure 5A). A plot of
the Lineweaver-Burk equation (eq 3) then yields Km and Vmax
1/ V0 ) (Km/ Vmax)/ [S] + 1/ Vmax
(3)
Figure 5. Analysis of data obtained under homogeneous reaction
conditions for the HRP-catalyzed reaction between H2O2 and amplex
red to yield fluorescent resorufin. (A) Plot of normalized fluorescence
intensity vs time. H2O2 concentrations ranged from 0.1 to 5.0 µM.
Conditions: 0.01 unit/mL HRP plus 10 µM amplex red in 2.0 mL of
50 mM Tris-HCl buffer (pH 7.4). (B) Lineweaver-Burk plot of the
reciprocal initial reaction rates obtained from (A) vs the reciprocal of
the H2O2 concentration.
(Figure 5B). The results of this treatment yield Vmax ) 1.90 µM/
min and Km ) 1.55 µM. This value of Km is very close to the value
of K 0m(app) (Km ) 1.51 µM) determined using the continuous-flow
approach described earlier.
To demonstrate the generality of this approach, the â-Gal-
catalyzed hydrolysis of nonfluorescent RBG to
D-galactose and
using various RBG concentrations and flow rates, primary plots
of f[A0] against -ln(1 - f) were used to generate Km(app) values.
A secondary plot of Km(app) versus Q yields a K 0m(app) value of 409
µM (Figure 7C). This value compares reasonably well with that
determined in homogeneous solution (Km ) 363 µM) and with
literature values (Km ) 380 µM and 500 ( 200 µM).12,30
fluorescent resorufin was also investigated (Figure 6A). To validate
the microfluidic-based method for measuring enzyme kinetics, the
value of Km was first determined in homogeneous solution. Figure
6B shows plots of fluorescence intensity, arising from the
appearance of resorufin, as a function of time for RBG concentra-
tions ranging from 50 to 250 µM. The initial slopes from these
plots and the Lineweaver-Burk equation (Figure 6C) were used
to determine a Michaelis constant, Km, of 363 µM.
Next, the kinetics for RBG hydrolysis by â-Gal were evaluated
using the continuous-flow approach. RBG solutions varying in
concentration from 25 to 200 µM (buffer: 100 mM Tris-HCl, 2
mM KCl, and 0.1 mM MgCl2 at pH 7.8) were pumped into the
inlet of the microfluidic device at flow rates of 10, 30, 50, 70, and
100 nL/ min. As before, the reaction rates were determined by
capturing fluorescence images at the inlet and outlet of the
microreactor (Figure 7A) and then converting the corresponding
fluorescence intensities (determined from line scans, Figure 7B)
to RBG concentrations using a calibration curve constructed by
flowing resorufin only through the channel. From data obtained
SUMMARY AND CONCLUSIONS
We have described a continuous-flow method for measuring
enzyme kinetics in microfluidic systems. The approach is conve-
niently implemented using bead-immobilized enzymes contained
within microreactors having a liquid volume (the difference
between the total microreactor volume and the beads) of 0.45 nL.
It is possible to estimate the total number of enzyme molecules
required for the kinetic analysis by assuming each enzyme has a
projected area of 80 nm2, that the total surface area of the 275
beads in each microreactor is 2.1 × 105 µm2, and that the enzymes
are close-packed on the bead surface. This treatment suggests a
maximum of 3 × 109 enzyme molecules are required for the
analysis.
(29) Brahim, S.; Narinesingh, D.; Guiseppi-Elie, A. J. Mol. Catal. B 2 0 0 2 , 715,
1-12.
(30) Hofmann, J.; Sernetz, M. Anal. Chim. Acta 1 9 8 4 , 163, 67-72.
Analytical Chemistry, Vol. 75, No. 13, July 1, 2003 3165

Figure 7. (A) Fluorescence micrograph showing the enzymatic
production of fluorescent resorufin in the outlet stream of a microre-
actor packed with â-Gal-functionalized microbeads. Flow rate, 50 nL/
min. (B) Normalized fluorescence intensity line scans, obtained at
the location indicated by the dashed line in (A), at RBG concentrations
ranging from 25 to 200 µM. (C) Plot of Km(app), the apparent Michaelis
constant, vs Q.
Figure 6. (A) â-Gal-catalyzed reaction of nonfluorescent RBG to
yield D-galactose and fluorescent resorufin. (B) Normalized fluores-
cence intensity vs time for this reaction carried out in homogeneous
solution. RBG concentrations ranged from 50 to 250 µM. Condi-
tions: 0.1 unit/mL â-Gal in 2.0 mL of 100 mM Tris-HCl buffer (pH
7.8), 2 mM KCl, and 0.1 mM MgCl2. (C) Lineweaver-Burk plot of the
reciprocal initial reaction rate vs the reciprocal of the RBG concentra-
tion.
and after immobilization on the microbeads. Similar findings have
previously been reported for enzymes immobilized in fluid lipid
bilyaers.21
Because of the reduced cost, reagent economy, and increased
throughput associated with microfluidic-based assays, it is quite
likely that many analytical determinations presently carried out
in microtiter plates will be performed using microfluidic systems
in the not-too-distant future. This paradigm shift will necessitate
development of new standard methods for measuring kinetic and
thermodynamic properties of reactions. The results reported here
contribute to this important goal, and they are therefore likely to
The kinetic analysis relies on the Lilly-Hornby model, which
has been previously used to evaluate apparent kinetic parameters
in packed-bed enzyme reactors under steady-state conditions. The
Michaelis constants determined using the new approach reported
here and traditional methods based on the Lineweaver-Burk
relationship are indistinguishable within the estimated precision
of the methods. A significant outcome of this study was the finding
that the kinetics of both â-Gal and HRP were the same in solution
3166 Analytical Chemistry, Vol. 75, No. 13, July 1, 2003

be of value for clinical diagnostics, and drug discovery and
screening.
Cooperative Agreement NCC-1-02038. Some of the instrumentation
used to carry out this work was provided by the Center for
Integrated Microchemical Systems at Texas A&M University.
ACKNOWLEDGMENT
Financial support from the U.S. Army Medical Research &
Received for review February 17, 2003. Accepted April 8,
2003.
Material Command is gratefully acknowledged. We also acknowl-
edge the support of the Texas Institute for Intelligent Bio-Nano
Materials and Structures for Aerospace Vehicles, funded by NASA
AC034155B
Analytical Chemistry, Vol. 75, No. 13, July 1, 2003 3167