ORIGINAL PAPER
Compact, cost-efficient microfluidics-based
stopped-flow device
Regina Bleul & Marion Ritzi-Lehnert & Julian Höth & Nico Scharpfenecker & Ines Frese &
Dominik Düchs & Sabine Brunklaus & Thomas E. Hansen-Hagge &
Franz-Josef Meyer-Almes & Klaus S. Drese
Received: 14 July 2010 /Revised: 15 November 2010 /Accepted: 16 November 2010 /Published online: 30 November 2010
# Springer-Verlag 2010
Abstract Stopped-flow technology is frequently used to
monitor rapid (bio)chemical reactions with high temporal
resolution, e.g., in dynamic investigations of enzyme
reactions, protein interactions, or molecular transport
mechanisms. However, conventional stopped-flow devices
are often overly complex, voluminous, or costly. Moreover,
excessive amounts of sample are often wasted owing to
inefficient designs. To address these shortcomings, we
propose a stopped-flow system based on microfluidic design
principles. Our simple and cost-efficient approach offers
distinct advantages over existing technology. In particular,
the use of injection-molded disposable microfluidic chips
minimizes required sample volumes and associated costs,
simplifies handling, and prevents adverse cross-contamination
effects. The cost of the system developed is reduced by an
order of magnitude compared with the cost of commercial
systems. The system contains a high-precision valve system
for fluid control and features automated data acquisition
capability with high temporal resolution. Analyses with two
well-established reaction kinetics yielded a dead time of
approximately 8-9 ms.
Keywords Stopped flow. Microfluidics . Low sample
amounts . Short dead time
Introduction
First introduced in 1940 by Chance [1] and improved in 1964
by Gibson [2], stopped-flow technology is a highly versatile
and convenient method for monitoring fast reactions. It is
often used to study the dynamic interactions of biomolecules
or other macromolecules, where it can provide insights into
the molecular details of enzyme functions, protein–protein or
protein–DNA interactions, as well as molecular transport
mechanisms [3–5]. Another application is in the investigation
of protein folding kinetics [6]. Understanding these
kinds of processes is of growing importance not only for
basic research but also to the pharmaceutical industry. For
example, stopped-flow technology is key in elucidating
catalytic enzyme mechanisms and functions of specific
proteins in an organism. Reactant concentrations can be
monitored photometrically, and changes in enzyme conformation
during substrate binding or product release can be
tracked using fluorescent dyes. In some cases, changes in
spectral properties of the reactants can be exploited to
optically detect the binding process. A prerequisite, however,
is that the substances be fully mixed before the onset of
binding. Clearly, then, manual mixing is inadequate for
observing processes on the timescale of milliseconds; a more
efficient mixing method is needed [7]. Also, highly time
resolved data acquisition must be available as part of the
setup.
As a result of these stringent requirements, commercial
stopped-flow devices have tended to be complex in design,
incorporating various components (i.e., light source, amplifier
electronics, detection unit, mixing station) that in the aggregate
make these systems bulky, expensive to purchase and
maintain, and prone to mechanical maladjustments. Typical
equipment often contains two step-motor-driven syringes
which push the reactants (e.g., enzyme and substrate), at very
R. Bleul :M. Ritzi-Lehnert (*) :J. Höth :N. Scharpfenecker :
I. Frese :D. Düchs :S. Brunklaus :T. E. Hansen-Hagge :
K. S. Drese
Institut für Mikrotechnik Mainz GmbH (IMM),
Carl-Zeiss-Str. 18-20,
55129 Mainz, Germany
e-mail: ritzi@imm-mainz.de
R. Bleul :F.-J. Meyer-Almes
Hochschule Darmstadt,
Schnittspahnstr. 12,
64287 Darmstadt, Germany
Anal Bioanal Chem (2011) 399:1117–1125
DOI 10.1007/s00216-010-4446-5
high velocity and in one go, first through a mixing chamber
and then into the optical detection cell. The detection unit, in
turn, is triggered by a stop syringe. The processes within the
detection cell may be monitored via changes in absorption or
fluorescence intensity. Such monitoring can, however, only be
initiated after the so-called dead time has elapsed, which
denotes the time span between the first contact of the reactants
and the beginning of measurements in the detection region
which are usable for kinetics analysis. It is thus dependent on
both the mixing and the transport times. In effect, only
processes happening on timescales longer than the dead time
can be monitored. The dead time is determined mainly by the
fluid flow rate, the mixing efficiency, the volume traversed by
the fluid between the mixing chamber and the detection cell,
and the volume of the detection cell [8, 9].
The intricate setups of conventional stopped-flow systems
necessitate relatively high sample and reagent volumes during
the first operating cycle. In serial experiments conducted
without changing chemicals, however, sample consumption is
very low. Still, the volumes required in the first cycle are often
already prohibitive as samples and reagents are expensive or
hard to isolate. Volumes of up to 3 ml during setup and,
subsequently, of 1 ml per experiment are not uncommon.
Extensive test series probing various parameters, as are
commonly performed, thus require considerable reactant
quantities. Despite this high rate of sample consumption,
however, most of the volume is consumed rather wastefully in
the flushing and washing steps. Here is where our
microfluidics-based stopped-flow system provides substantial
improvements in simplicity and cost-efficiency: the use of
injection-molded disposable microfluidic chips reduces the
required sample volume to 20 μl even in the first experiment
as flushing is no longer necessary; sample costs are hence
lowered significantly; handling is simplified; and adverse
cross-contamination effects are prevented.
The chip layout and material choice allow for the use of
narrow-bandwidth light-emitting diodes (LED) as light
sources in the UV–vis spectrum instead of conventionally
used xenon and mercury arc lamps. These UV and visible
LEDs can easily be replaced with ones covering different
wavelengths as the case may demand, thus broadening the
applicability of our approach. Furthermore, UV–vis-sensitized
photodiodes covering a wide range of wavelengths are
employed instead of photomultipliers for detection.
In our case, a uniform illumination of the flow channel
over a length of 10 mm is achieved with integrated on-chip
mirrors that reflect light toward the detection cell and the
detector. Coupling the photodiode with simple but effective
amplifier electronics offers excellent sensitivity and resolution
over the spectral range relevant in most biological applications
(i.e., 250-1,000 nm).
The overall cost of the stopped-flow device at current
component prices, including costs for housing, electronics,
and optical elements, is in the low four-digit euro range,
which is roughly an order of magnitude lower than that of
commercially available solutions. Moreover, economies-ofscale
effects are expected to further lower the per-unit cost
at higher production volumes.
The performance is studied using two chemical reactions
as test cases:
1. Reaction I: the reduction of 2,6-dichlorophenolindophenol
(DCIP) by ascorbic acid according to Tonomura et al. [8]
2. Reaction II: the fluorescence quenching of Nacetyltryptophanamide
(NATA) by N-bromosuccinimide
(NBS) according to Peterman [9]
The functionality of our system is evaluated by reaction
III: labeling histone deacetylase like amidohydrolase
(HDAH) from Bordetella with fluorescent dyes [19]. This
bacterial amidohydrolase is structurally highly homologous
to, and thus serves as an easily produced proxy for, the
human histone deacetylases currently under investigation as
possible targets for cancer therapy. Because of this high
homology, the transferability of drug screening results is
expected in many cases.
Materials and methods
Polymer chips
Poly(methyl methacrylate) (PMMA; Plexiglas® 7N) was
chosen as the polymer chip material owing to its high degree
of transparency in the UV range and low autofluorescence,
which allows dynamic protein investigations at a wavelength
of 280 nm (the intrinsic fluorescence wavelength of trypto-
phan).1
Structures for fluid transport and mirrors for light
reflection were machined into injection-molded chip blanks
by precision milling and laser ablation technologies. The
outer chip dimensions are 43 mm×64 mm. The channels
for reactant inlets are 1 mm×1 mm, having a semicircular
shape at the bottom (radius 0.5 mm). The T junction
has a depth of 0.4 mm and a width of 0.5 mm. The
length of the tapered channel is 9.5 mm and the diameter at
the connection of the detection cell is 1 mm. The detection cell
is 10 mm long and 1 mm in diameter. The volume of the
complete structure is 64.5 μl, where the volume from the T
junction to the end of the detection cell is 16.5 μl (Figs. 2, 3).
The structured chip is bonded with microencapsulated foil
(40 μm, polyolefin, HJ Bioanalytik, Mönchengladbach). To
enhance the wettability of the channel surfaces and suppress
unspecific adsorption processes, the surfaces were processed
1
A PMMA 7 N layer of 2 mm blocks approximately 50% of the light
at 280 nm. The optical path through the chip material has a length of
1 mm before plus 1 mm after the detection cell.
1118 R. Bleul et al.
with ethylenediamine or ethanolamine as described by Stein
et al. [19].
Instrument setup
For the electronics, controlled high-precision magnetic
valves purpose-made for miniaturized instruments (The Lee
Company, Westbrook, CT, USA) were used. They combine the
advantages of a very fast response time (less than 1 ms), a
small dead volume (a few microliters), and a low operating
voltage (12 V). Valves resistant to pressures up to 120 psi
(approximately 8 bar) are available. Since these valves are
prone to contamination with small particles, it is recommended
that the liquids be filtered before induction (preferably using
pores with diameters of 10 μm). Syringe pumps are used to
load the samples and to pump them without pulsation (cetoni,
Korbußen/Gera, Germany).
The detection unit consists of a LED for excitation (Roithner
Lasertechnik, Vienna, Austria) and photodiodes for absorption
and fluorescence detection (Silicon Sensor, Berlin, Germany).
The photodiodes are equipped with a cutoff or band-pass
interference filter for wavelength selection. Alternatively,
absorbing filters made of a bulk polymer such as polycarbonate
(Makrolon® UV 2099, 2 mm, Röhm, Darmstadt, Germany) or
polystyrene (1 mm, Greiner Bio-One, Solingen, Germany)
could be used to separate the excitation light from the
fluorescence signal if their peak wavelengths are sufficiently
distant from each other (e.g., deep-UV excitation at 280 nm
with UV fluorescence at 335 nm, or UV excitation at 385 nm
and visible fluorescence at 480 nm).
The beam propagation inside the chip prevents direct
exposure of the fluorescence detector to the excitation beam.
Since the photoelectric current of the diodes is in the picoampere
to microampere range, a transimpedance amplifier (sglux
SolGel Technologies, Berlin, Germany) is used for signal
enhancement and to transform current signals to voltage signals,
the latter with amplification factors of 107
V/A (off-line) or
1010
V/A (in-line) and output signals not exceeding ±4 V. The
high amplification factors used require the system to be
shielded electrically with an aluminum cover, which also
serves as an optical shield blocking ambient light.
The device is connected to a computer via a multifunctional
data acquisition board (16-bit; National Instruments, Austin,
TX, USA) providing analog input ports for data from the
optical sensor and digital output ports enabling valve control
in the millisecond range. The data are collected at selectable
sample rates of up to 80,000 samples per second for a
specified total measurement time. The instrument is controlled
with LabVIEW 8.6 (National Instruments). The setup realized
is shown in Fig. 1.
Figure 2 shows a schematic diagram of the microchipbased
stopped-flow device. Solutions A and B are pushed
into the chip using syringe pumps up to stop valve V-6,
thus filling the system with liquid free of air bubbles
throughout the microfluidic channel geometry. After
switching valves V-4 and V-5, valve V-1 is opened for a
predetermined time (e.g., 20 ms) to drive both solutions
into the detection cell by virtue of a gradient in air pressure
of 2 bar. The flow is actively stopped by simultaneously
closing valves V-1 and V-6. Data acquisition starts upon
pumping to monitor the complete process, and all measured
data are stored for subsequent analysis. Once the measurement
is complete, first valve V-6 is opened, then valve V-2, to
ventilate the system. An additional valve and pump are
integrated into the system so it can be washed after each
experiment. To do so, wash buffer is pumped through the
system from valve V-3, which has to be opened for this task.
Mixing
As mentioned already, fast mixing is a major requirement in
highly time resolved measurements. Various mixers with
mixing times down to the millisecond range are discussed
in the literature [21–25]. Commercially available stoppedflow
instruments most often contain ball mixers [26]
utilizing the turbulent wake which occurs downstream from
a spherical surface. Since mixing is ultimately a diffusion
process on microscopic length scales, mixers generally aim
to increase the contact area of the reactants to achieve faster
mixing [21–25]. This is often achieved by subjecting the
flow to turbulent flow conditions.
Falk and Commenge [25] recently demonstrated that the
main factor determining mixing times is the pressure drop
across the mixer. They further showed that the mixing time is
governed by a common empirical scaling behavior with
respect to the pressure drop across a wide array of mixer
geometries and flow rates. The task of finding an appropriate
mixer thus reduces to selecting a geometry that will produce
the required pressure drop at the pertinent flow rate. Using
this result, which holds in both the laminar and the turbulent
flow regimes, we conclude that a basic T mixer geometry
[27] suffices to obtain the required mixing performance.
Adjusting the pressure drop via variations in the spatial
dimensions of the mixing section allows this mixer to easily
fit into the overall layout of our system. At the junction of
the T structure, the two solutions collide and merge at high
velocities. The reactants are mixed by the induced vortices,
and the reaction starts within the short mixing channel [28].
Upon leaving the mixing channel, the mixture is propelled
directly into the optical detection cell.
A flow rate of 3.6 ml/s was determined by simulating
fluid velocity fields within the chip at 2-bar overpressure
using the computational fluid dynamics program Flow 3D
(Flow Science, Santa Fe, NM, USA). The velocity is
highest at the beginning of the mixing section, being
approximately 20 m/s (Fig. 3). For aqueous solutions, the
Compact, cost-efficient microfluidics-based stopped-flow device 1119
calculated Reynolds number is around 9,000, indicating
turbulent flow conditions (turbulence is generally found
above Reynolds numbers of approximately 2,300).
Using Falk and Commenge’s results, the residence time
of the fluids over the T junction and the tapered section (a
few milliseconds) is approximately 40% higher than the
required mixing time for the geometry employed. Moreover,
this ratio remains roughly constant throughout the
range of typical pressures thereby ensuring satisfactory
levels of mixing. Commercially available high-performance
stopped-flow systems, however, offer flow rates of 13 ml/s
and dead times less than 1.5 ms (Table 1).
The flow may be accelerated by increasing the applied
pressure. As a result, turbulences will tend to be amplified,
leading to faster mixing and shorter dead times (as defined
above).
Dead time determination by absorption and fluorescence
measurements—reactions I and II
The dead time of the stopped-flow device is defined as the time
interval from the onset of the reaction of interest to the time
when it can first be monitored. It depends mainly on the mixing
time, the flow rate, and the dead volume and can be determined
from signals (e.g., absorption or fluorescence) measured in
simple first-order reactions [8, 9]. To examine the dead time
of our device, we measured reaction I (by absorption) and
reaction II (by fluorescence) as described in “Introduction.”
In both cases, a signal (absorption or fluorescence) is
obtained which varies as the chemical reaction progresses.
Although the reaction starts immediately upon first contact of
the reactants, this initially happens outside the detection zone.
Moreover, to obtain reliable data, valves V-1 and V-6 are
closed after the initial transport time has elapsed and the
fluid volume previously present in the detection zone has
been flushed out. After the fluid flow has been stopped, it
still takes approximately 10 ms before data points usable
for kinetics analyses are obtained (Fig. 4).
We therefore discard all data taken less than 10 ms
after the simultaneous closing of valves V-1 and V-6
and shift the time axis to this point in time, denoting it
by t=0. The resulting graphs are characterized by a quick
ascent at short times followed by an equilibrium state at
long times, as one would expect from a saturating
Fig. 2 The microchip-based stopped-flow device
a b
mirrors
detection cell
Fig. 1 a Laboratory setup of
the microfluidic chip-based
stopped-flow device [poly
(methyl methacrylate) (PMMA)
chip dimensions 64 mm×
43 mm]. b Close-up of detection
cell with surrounding integrated
45° surfaces enabling internal
reflection of excitation light
(small mirrors) as well as
fluorescence (larger lateral
mirrors) inside the PMMA
chip
1120 R. Bleul et al.
reaction (also see Fig. 5a). An appropriate fit for this
behavior is given by Eq. 1:
a þ bt þ ceÀkt
; ð1Þ
where the term bt represents the influence of drift in the
measuring device. The fit parameters a, b, c, and k are
calculated with the solver integrated in Microsoft Excel.
For better statistics, this is not done on results from a
single experiment. Rather, we run the same experiment
several times and normalize the resulting data sets by
multiplying them by gauge factors such that the medians
of the last ten data points of each graph coincide. Then
we carry out the fit on the averaged signal plot.
For each set of experiments at different reactant concentrations,
we repeat the procedure just described. The rate
constant, k, of the reaction is obtained for each concentration
directly as part of the respective fit. To determine the dead
time, we proceed as follows. First, the median-normalized and
averaged signal plots for the different reactant concentrations
(as shown, e.g., in Fig. 5b) are shifted relative to each other
such that the endpoints of their fits coincide. The objective
here is to gauge all graphs to a common asymptote
representing the saturation stage of the reaction. It is pertinent
that the drift parameter, b, varied only very slightly (typically
less than 2%) across fits for different concentrations, justifying
our normalization procedure.
The point at t<0 where the graphs intersect marks the
lower end of the dead time interval. The upper end is given
by t=0. Following Peterman [9], we plot the logarithm of
the graphs after subtracting their long-time limits (thereby
“linearizing” them; see, e.g., Fig. 5c) to better discern their
common crossing point. The dead time is then given by the
distance on the time axis between this point and t=0.
HDAH expression, isolation, and labeling
with fluorescamine–reaction III
HDAH was expressed in, and purified from, bacterial strain
XL1-Blue-FB188-HDAH. The strain and protocols were
kindly provided by C. Hildmann (TU Ilmenau, Ilmenau,
Table 1 Dead times of commercial systems
Company/product Dead time (ms) Reference
Applied Photophysics SX 18MV ≈2 [10]
Applied Photophysics SX MV 2 (Fluo),
8 (CD)
[11]
Applied Photophysics SPF-17 15 [12]
AVIV Instruments stopped-flow tower interface with AVIV 202 SF spectrometer 5 [13]
BioLogic SFM-4 15 [14]
BioLogic SFM-20 <1.5 [15]
TgK Scientific HPSF-56 <10 [16]
Unisoku RSP-1000 (Japan) 3 [17]
Unisoku type USP-539 15 [18]
velocity magnitude in m/s
yinm
x in m
detection cell
T mixer
Re=9000
Fig. 3 Snapshot of velocity
magnitude distribution of the
microfluidic T mixer structure
with an adjacent tapered channel
and detection cell obtained by
transient computational fluid
dynamics simulation using
Flow3D. Dark blue represents
open regions in the chip such as
mirrors and bore holes. T mixer
dimensions 0.5-mm×0.4-mm
channel diameter, 3-mm outlet
length, 9.5-mm-long tapered
channel
Compact, cost-efficient microfluidics-based stopped-flow device 1121
Germany). The HDAH concentration was determined using a
bicinchoninic acid protein determination kit (Interchim) as
prescribed by the manufacturer. When excited at 390 nm,
fluorescamine (4-phenylspiro-[furan-2(3H),1-phthalane]-3,3′dione)
exhibits strong fluorescence at 475-490 nm when
bound to proteins. For the reaction of 3 μM HDAH in 0.2 M
borate, a buffer with a pH of 8.6 was mixed 1:1 with 1-4 mM
fluorescamine in dimethyl sulfoxide. For optical detection, a
385-nm LED was used. The amplifier gain was set to
5·107
V/A.
The data from reaction III were fitted to the
expression a + bt + c exp(-kt) in the same fashion as for
reactions I and II to obtain reaction rate constants, k. Since
the dead time is not a critical parameter for this analysis, it
was not determined again here. The signal plots used
(Fig. 7) show clearly undisturbed data, justifying our
analysis procedure.
Results and discussion
Dead time determination by absorption
measurements—reaction I
The first reaction used to determine the dead time of the
system was the reduction of DCIP by ascorbic acid (Fig. 5).
Data were acquired at a sample rate of 10,000 s-1
(individual measurements, no integration was performed)
starting 10 ms after the closing of stop valve V-6. Analyses
of seven independent measurements revealed an average
rate constant, k, of (94±13)s-1
. After averaging and data
fitting, we obtained a rate constant, k, of 91.1 s-1
(Fig. 5a).
The reaction speed can be influenced by varying the
concentration, yielding half-lives shorter than 1 ms. For the
reaction, 5-20 mM ascorbic acid in 0.2 M NaCl at pH 4 was
mixed 1:1 with 140 μM DCIP. For the optical detection, a
525-nm LED and a detector with an interference filter
(536 nm, band width 40 nm) were used. The amplifier gain
was set to 109
V/A.
Repeating the same experiments with different ascorbic
acid concentrations (2.5-10 mM) reveals a nearly linear
dependence of the rate constant on the ascorbic acid
concentration (inset in Fig. 5b). The dead time is obtained
as described above and amounts to approximately 9 ms,
which is competitive with conventional stopped-flow
systems, e.g., from TgK Scientific (Bradford-on-Avon,
UK), whose dead times are specified as 8-10 ms.
Rate constants for the DCIP reduction by ascorbic acid,
however, are difficult to compare since for pH 4 Tonomura
et al. [8] only reported data for a higher ascorbic acid
concentration (20 mM). Moreover, between pH 4 and pH 5,
the rate constants are rather sensitive to the pH of the
reaction mixture (pH 4.07, k=838 s-1
; pH 4.91, k=224 s-1
[8]). Yet, the rate constant of 149 s-1
(Fig. 5) determined
with our stopped-flow device for 10 mM ascorbic acid lies
in the anticipated range, since a linear extrapolation of pHdependent
data given in [8] to lower concentrations results
in k values of 419 and 112 s-1
for pH 4.07 and pH 4.91,
respectively.
Dead time determination by fluorescence
measurements–reaction II
Another standard reaction for the determination of the dead time
is the pseudo-first-order quenching reaction of NATA by NBS,
which exhibits rate constants, k, ranging from 70 to 155 s-1
[9].
Experiments were conducted using 10 μM NATA (SigmaAldrich)
mixed 1:1 with 200-400 μM NBS (Sigma-Aldrich),
leading to total NBS concentrations of 100-200 μM. The
excitation wavelength was 280 nm, and the emission was
measured above 300 nm using a polystyrene filter. All
Fig. 4 Signals recorded after closing the valves: a absorption in
reaction I at several ascorbic acid concentrations, and b fluorescence
intensity in reaction II at several N-bromosuccinimide (NBS) concentrations.
The time axis is shifted to the start of the first usable data for
kinetics analysis (t=0, dashed line)
1122 R. Bleul et al.
solutions were freshly made in degassed 50 mM phosphate
buffer, with a pH of 7 in a nitrogen atmosphere. For the optical
detection, the LED was combined with a polystyrene
absorption filter. Being able to conduct measurements in the
UV range is of particular interest in investigations involving
the intrinsic fluorescence present in proteins.
An analysis of four independent measurements for each
NBS concentration is shown in Fig. 6. It can be seen that
the data are approximated well by exponentials (the drift
parameters, b, are within a few percent of each other and so
can be neglected in evaluating the dead time, as per the
earlier discussion). A dead time plot was obtained in the
same fashion as for the absorption experiments, again
yielding a dead time estimate of approximately 8 ms and
thus corroborating the earlier result. The first-order rate
coefficient, k, determined for the quenching of NATA
Fig. 6 Quenching of N-acetyltryptophanamide (NATA; 5 μM) by
NBS, pH 7, 23 °C: a Average of four independent measurements of
fluorescence intensity, F, for each NBS concentration. b Determination
of dead time (dashed line at t=–8 ms) and first-order rate coefficient, k,
for the quenching of NATA fluorescence by NBS with the present
stopped-flow system in comparison with published data [9]
Fig. 5 Reduction of 2,6-dichlorophenolindophenol (DICP) by ascorbic
acid. a Individual absorption measurements with 70 μM DCIP+5 mM
ascorbic acid, pH 4, 22 °C. b Average of seven independent measurements
for each ascorbic acid concentration (70 μM DCIP). The
inset shows the linear dependence (R2
=0.9611) of the rate constant,
k, on ascorbic acid concentration, c. c Dead time plot of linearized
curves from b. The resulting dead time is approximately 9 ms
(dashed line)
Compact, cost-efficient microfluidics-based stopped-flow device 1123
fluorescence by NBS with the present stopped-flow system
is comparable to published data [9].
Functionality evaluation using HDAH–reaction III
Since the system seemed capable of determining reaction
kinetics reproducibly and in accordance with published
data, additional experiments were done to evaluate its
functionality. As a test case for a two-step reaction, labeling
of HDAH with fluorescamine was examined, where the
following mechanism was proposed by Stein et al.2
:
F þ A Ð
k1
kÀ1
F Á A½ Š !
k2
P: ð2Þ
F represents fluorescamine, A stands for HDAC, [F∙A]
denotes an intermediate complex, and P is the fluorescent
labeled product. From the experimental results shown in
Fig. 7, an apparent pseudo-first-order reaction rate, k, can be
determined. Following the proposed reaction mechanism, k1/k-1
and k2 are calculated utilizing the linear relation
1
k
¼
1
k2
þ
kÀ1
k1k2
1
F½ Š
: ð3Þ
The equilibrium constant for intermediate complex formation,
k1/k-1, as obtained from Eq. 3 is 0.9 mM-1
, whereas in [19] a
corresponding k1/k-1 of 0.2 mM-1
was reported for a similar
reaction of fluorescamine with a primary amine (alanine).
The rate constant k2≈78 s-1
of the subsequent irreversible
reaction step, however, is fairly comparable to k2≈100 s-1
given in [19]. Therefore, both values confirm the suitability
of our stopped-flow device for analysis of two-step reaction
kinetics.
Conclusion and outlook
A microchip-based automated stopped-flow device has
been presented whose small size and significantly reduced
reagent consumption represent an important advance for the
measurement of highly time resolved reaction kinetics. The
present system now puts applications previously not
considered feasible, owing either to prohibitive costs or
insufficient amounts of available reagent volumes, within
the realm of possibility. Based on a microfluidic polymer
chip and a compact custom-made instrument, the system is
flexible in application, easy to handle, and cost-efficient in
both acquisition and operation. The overall cost of the
system presented here (low four-digit euro range) is roughly
an order of magnitude lower than that of commercially
available systems. Economies-of-scale effects are expected
to further reduce per-unit costs and thus open up a path to
providing low-cost solutions to end users.
Allowing linear absorption measurements up to about
A490 =1.5 and a lower detection limit of fluorescein
(excitation wavelength, 490 nm; emission wavelength
530 nm; amplification 107
V/A) at 50 nM, it is as efficient
as most conventional photometers (the most sensitive
fluorescence spectrometers have detection limits down to
5 nM fluorescein [29]). The integration of a stop valve
helps make the fluid control more robust. The possibility to
wash the system after each experiment allows for consecutive
series of measurements on the same chip.
To validate the performance and to determine the dead
time of the microchip-based stopped-flow device, three
different reactions were investigated. Also, measurements
in the UV range can be conducted, e.g., for investigations
of the intrinsic fluorescence of proteins. Our investigations
Fig. 7 Labeling of histone deacetylase like amidohydrolase (HDAH)
with fluorescamine. a Fluorescence intensity measured for 3 μM
HDAH mixed 1:1 with fluorescamine in dimethyl sulfoxide at 22 °C;
average of four independent measurements for each fluorescamine
concentration. b Natural logarithm of the difference of the maximum
observed fluorescence intensity and time-dependent intensity, plotted
against time. The values determined for k are shown
2
The reaction was set up following a protocol described in [19] for
the labeling of alanine.
1124 R. Bleul et al.
showed agreement with data published in the literature for
all three test reactions. In particular, two reactions showed
dead times of approximately 8-9 ms, which is competitive
with conventional stopped-flow systems. In the third
reaction, kinetic parameters were reproduced consistent
with the published reaction scheme.
Although our system is already competitive with
commercially available stopped-flow systems, a further
reduction in dead times to approximately 0.25 ms will be
required to approach the performance level of today’s bestperforming
systems. Another goal is the further reduction
of sample volumes. In this pursuit, reducing the channel
length before the mixing structure and behind the detection
cell may prove advantageous. Therefore, we envisage the
integration of fast switchable valves on the chip. Despite
the turbulent flow conditions prevalent in our system, the
mixing efficiency can and must be enhanced. Introducing
structures to that effect should shorten mixing times, which
would be desirable especially for cases of asymmetric
reagent volumes. Application of higher pressures will
reduce the dead time further but may come at the price of
reduced reproducibility due to higher turbulence. By careful
design of the mixing structure, it may be possible to reduce
dead times below the 1-ms mark.
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