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ntegration of microfluidics and FT-IR microscopy for label-freetudy of enzyme kinetics
vgeny Polshina, Bert Verbruggena, Daan Wittersa, Bert Selsb, Dirk De Vosb,art Nicolaï a, Jeroen Lammertyna,∗
KU Leuven – University of Leuven, Department of BIOSYST – MeBioS, Willem De Croylaan 42, B-3001 Leuven, Belgium1
KU Leuven – University of Leuven, Center for Surface Chemistry and Catalysis, Kasteelpark Arenberg 23, B-3001 Leuven, Belgium
r t i c l e i n f o
rticle history:eceived 19 October 2013eceived in revised form 27 January 2014ccepted 28 January 2014vailable online 5 February 2014
eywords:
a b s t r a c t
In this article we report on the integration of microfluidics with FT-IR microscopy for the label-free studyof enzyme kinetics. The IR compatible microfluidic chip was fabricated by standard photolithographyprocesses using a photopatternable PDMS and infrared transparent materials (Si and CaF2). Chip char-acterization was performed with an imaging focal plane array (FPA) detector. The enzymatic oxidationof glucose catalyzed by glucose oxidase, which served as a model system, was monitored on-chip in realtime in a label-free manner using FT-IR microscopy. The reference FT-IR measurements were carried out
T-IR microscopyttenuated total reflectionhemical imagingicrofluidicsicrofabrication
nzyme kinetics
using the attenuated total reflection (ATR) accessory. Michaelis–Menten parameters for glucose-oxidasewere estimated from the spectral measurements both on-chip and off-chip. The proposed microfluidicapproach for enzyme reaction monitoring serves as a novel strategy for FT-IR microscopy allowing forminimal reaction volumes, measurement automation and flexibility in terms of spatial, spectral and tem-poral data acquisition and offers new opportunities in kinetics studies of various bio(chemical) reactions.
There is a clear trend in the analytical domain toward miniatur-zation, simplification, automation and parallelization of analyses.
icrofluidic technology downscales the sample volumes andllows the integration of unit operations on lab-on-a-chip plat-orms in order to reduce the analysis time and cost, increasehroughput and achieve higher sensitivity levels [1]. Detectionf chemical or biological species in microfluidic platforms isommonly achieved by optical methods, such as fluorescencepectroscopy, owing to its superior sensitivity down to singleolecule detection [2]. However, the disadvantage of the fluores-
ence method is the need for additional reaction steps, i.e., labelingolecules with fluorescent tags, which in turn may potentially
ffect the behavior of the system being investigated. Therefore,nterest has grown in integrating label-free detection techniques
ith microfluidics platforms [3]. One such label-free techniqueith very broad applicability is Fourier Transform Infrared
FT-IR) microscopy, which is an information-rich research tool that
provides not only spectral information, but also spatially resolveddata about the distribution of the sample constituents at any mid-infrared (MIR) wavelength [4]. FT-IR microscopy allows to work intransmission, reflection and attenuated total reflection (ATR) samp-ling modes using single point, mapping or hyperspectral imagingoperational principles. This versatile method has opened the doorto new applications for in situ microspectroscopic mapping andimaging of various samples. Furthermore, FT-IR microscopy hasrecently been applied as a novel detection concept in microflu-idics [5–9]. Among many proposed approaches one of the mostinteresting is the micromixer designed for time-resolved FT-IRmeasurements [9], which can be considered as a benchmark in thisfield. It allows monitoring fast chemical reactions using FT-IR detec-tion in the low millisecond time regime, but requires the use of aseries of complex semiconductor processing steps for device fabri-cation, thus limiting its adoption by other researchers. In this articlewe describe a less complex and sophisticated fabrication approachsuited for slower reactions.
Both FT-IR spectrometry and microscopy are known to be excel-lent methods to study time-transient processes in real-time in a
label-free manner [10]. One of the examples is the investigationof different chemical and biochemical processes, such as enzy-matic reactions [11]. Enzymes are fundamental to life and arewidely used in many bio-assays and biological processes due to
heir exceptional catalytic activity. Therefore, the investigation ofnzymatic reactions is of great interest in many contexts. Mostften, the enzymatic reactions cannot be followed directly with UVr vis spectroscopy and usually it is required to couple the reac-ion of interest to auxiliary enzymatic reactions to produce easilyetectable species, such as NAD(P)H or chromophores [11]. How-ver, in such linked enzyme systems additional components mayntroduce disturbance and alter the kinetic behavior of the mainnzymatic reaction [12]. The application of FT-IR spectroscopyliminates the need for secondary reactions and can be used forirect monitoring of the enzymatic activity owing to molecular spe-ific information of the substrate and reaction products providedy their unique MIR spectra. Literature reports several examples ofhe monitoring of different enzymatic reactions on the basis of FT-R spectroscopy [11,12]. However, there are no reports on the use of
microfluidic platform for kinetic studies of enzymatic reactionsy FT-IR microscopy. The integration of microfluidics, as a novelample presentation strategy, with FT-IR microscopy offers newpportunities in obtaining spatially and temporally resolved chem-cal information on dynamic biomolecular processes. The objectivef this research is to design and fabricate a prototype microfluidicevice for integration with FT-IR microscopy and to demonstratehe potential of this integrated technology for measuring enzymaticeaction kinetics.
aterials and methods
abrication of the IR microfluidic chip
The design of the microfluidic chip was based on a simple-shaped channel geometry with 2 inlets and 1 outlet. All 3 chan-els were 15.3 mm long and 250 �m wide. The microfluidic chiponsisted of two parallel wafers of IR transparent materials: Si andaF2 with a gap size of 10 �m (channel’s depth) to reduce the waterbsorption barrier. A Si wafer (Siliciumbearbeitung A. Holm, Tann,ermany, 99.9999% purity) with a thickness of 675 �m was useds a bottom substrate of the chip. The CaF2 wafer (S.A.F.I.R, Knokke,elgium, thickness of 2 mm) was used as a top cover. In additiono IR transparency, CaF2 is also transparent in the visible spectralange allowing for visualization of the channels and alignment withhe IR beam of the FT-IR microscope. Both wafers were 50.8 mm iniameter.
Prior to the IR chip fabrication, the access holes (diameter.6 mm) for sample delivery were drilled in CaF2 wafer with aiamond bit at fast rotating speeds (6500 rpm). The IR chip wasabricated based on a standard photolithography process [13]. First,oth the Si wafer and CaF2 wafer were thoroughly cleaned beforehe fabrication. The Si wafer was cleaned in piranha solution (65%f H2SO4 + 35% of H2O2) for 15 min, rinsed with water and driedn a nitrogen flow, followed by heating on a hot plate for 2 mint 110 ◦C to remove any residue of water. The CaF2 wafer wasleaned in acetone for 10 min, followed by isopropanol for another0 min and air dried. Second, a Ti primer (MicroChemicals GmbH,lm, Germany) solution was spin-coated at 4000 rpm for 1 minn one side of the CaF2 wafer and oxidized afterwards on a hotlate at 110 ◦C for 10 min to form a 1 nm TiOx layer. This proce-ure resulted in a better adhesion between the PDMS layer andaF2 wafer during the bonding procedure. Third, the Si wafer waspin-coated with photopatternable PDMS (‘WL-5351’, Dow Corn-ng Corporation, Midland, MI, USA) at 1500 rpm (10 �m thick) andoft baked at 110 ◦C for 2 min. Subsequently, the photomask of theesigned channel pattern was brought in close proximity to the
DMS layer and exposed to UV radiation (Karl Suss MJB55 maskligner, Suss MicroTec, Germany) for 12 min. During the photo-ctivation process the channel pattern was transferred onto theegative photoresist (WL-5351). After the UV exposure the wafer
tors B 196 (2014) 175–182
was soft baked at 150 ◦C for 2 min and the designed channel patternwas developed in a negative resist developer (NRD). The standardprocedure of spin-coating WL-5351 normally requires a hard bakeat 250 ◦C for 30 min after the development to cross-link PDMS to theSi wafer. However, this step was skipped as the developed struc-tures were still sticky after the development process and thereforeit was possible to bond both Si and CaF2 wafer at room temperaturewithout any activation step (such as oxygen plasma activation).Finally, the Si and CaF2 wafers were sealed by the cold bondingprocedure for 4 h [7,8]. The scheme of the fabrication process isdepicted in Fig. 1.
When the fabrication process was finished, Nanoports (Achromnv, Machelen, Belgium) were aligned with inlet holes and attachedto the IR chip. The Nanoports were connected to a precision syringepump (PHD 2000, Harvard Apparatus) through polyethylene tubingused for sample delivery into the channel network. The microfluidicchip was held in a holder made from PMMA using the CO2 lasercutter (Trotec Ltd., Wels, Austria).
Sample preparation
For the enzyme kinetic study phosphate buffered saline (PBS)buffer (20 mM, pH 7.2), glucose oxidase (GOX) from Aspergillus niger(type X-S, lyophilized powder) and d-glucose were supplied bySigma Aldrich (Steinheim, Germany). The activity of the GOX was147,900 units/g solid. A 45 �M solution of GOX was made in 20 mMPBS buffer. The samples were stored at −20 ◦C and were thawedand warmed to room temperature (22 ± 1 ◦C) immediately beforethe measurements with ATR/FT-IR or on the chip. Fluoro-carbonoil FC-40 (3 M, St. Paul, USA) was used for generation of laminarflow that is immiscible with water flow inside the microchannels.All samples were filtered through a filter with 0.45 �m pore size(Millipore, Billerica, USA) prior to injection into the chip.
were performed on a Tensor 27 FT-IR spectrometer (Bruker OptikGmbH, Karlsruhe, Germany) equipped with a silicon carbide MIRsource, a Michelson interferometer with a potassium-bromidebeam-splitter and a liquid nitrogen cooled single point MCT detec-tor (Kolmar Technologies Inc., Newburyport, MA, USA). The FT-IRdifference spectroscopy approach was used for following theenzyme kinetics [12]. First, 25 �L of glucose (concentration range of12.5–300 mM) and 25 �L of GOX solution in 20 mM PBS buffer weremanually mixed in the Eppendorf vial (1.5 mL), vortexed and placedon the ATR accessory. Immediately after that the background spec-trum and the repeated sample spectra were collected with a timeresolution of 10 s for 40 min. By doing so, the main spectral contrib-utions of the substrate and the enzyme were subtracted from theresulting difference spectra, revealing the formation of the productabsorption bands. The ATR/FT-IR measurements were performedusing 32 co-added scans at a resolution of 8 cm−1.
On-chip transmission FT-IR microscopyThe transmission FT-IR microscopy measurements were car-
ried out on a Cary 620 FT-IR microscope coupled to a Cary 670FT-IR spectrometer (both Agilent Technologies Inc., Santa Clara,USA). The FT-IR microscope was equipped with a cryogenicallycooled FPA MCT camera and a single point MCT detector. The FPAdetector had 4096 pixels (40 �m2 each) with an imaging area of
350 �m × 350 �m with an option of field expanding optics thatquadruple the area of the analysis to 700 �m × 700 �m.
For generation of FT-IR Images 32 scans at spectral resolu-tion of 8 cm−1 were acquired for both the background and sample
E. Polshin et al. / Sensors and Actuators B 196 (2014) 175–182 177
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ig. 1. Schematic diagram of the fabrication process: (A) spin coating of PDMS ontDMS in NRD; and (D) sealing with CaF2 wafer.
easurements. The background measurements were created usinghe positions on the chip that were free of the PDMS layer. ThePA imaging detector was used for capturing the chemical images,hile a single point narrowband MCT detector (250 �m × 250 �m)as used for enzyme kinetic studies on the chip. The glucose solu-
ion was introduced through a channel from the right side of thehip and the glucose oxidase solution was pumped through a chan-el from the left side of the chip. The flow speed was 1 �L/min.oth streams were joined in the middle of the chip at a T-junction
orming a laminar flow, which was moved along the third chan-el toward the detection point (Fig. 2). A similar approach as
or ATR/FT-IR was employed here. First, the background scan of
he laminar flow of the two reagents was taken. Immediatelyfter that, the flow was stopped and the kinetic sample spectra32 scans) were collected every 10 s for 40 min at a resolutionf 8 cm−1.
ig. 2. (A) IR microfluidic chip used for studying enzyme kinetics by FT-IR microscopy andmaging FPA.
afer and soft baking; (B) UV exposure through the photomask; (C) development of
Data analysis
The collected data were analyzed using spectroscopic softwareOPUS (Bruker Optik GmbH, Karlsruhe, Germany, version 6.5) andResolutionPro (Agilent Technologies Inc., Santa Clara, USA, version5.2.0). The kinetics of the reaction were determined by nonlin-ear regression using the Michaelis–Menten equation in GraphPadPrism software (San Diego, USA, version 6).
Results and discussion
FT-IR chemical imaging
First, the IR microfluidic chip was characterized with the FPAimaging detector. A vis image of the T-junction in the middle of theIR chip is shown in Fig. 3(A). The corresponding FT-IR images were
(B) enlarged area shows the field of view of both detectors – single point MCT and
178 E. Polshin et al. / Sensors and Actuators B 196 (2014) 175–182
Fig. 3. Vis (A) and FT-IR images of the IR chip based on the absorption of PDMS at 2962 cm−1 (B) and 1260 cm−1 (C). Scale bars represent 100 �m. The MIR absorption spectrumof PDMS (D) was extracted from the upper right corner (at the cross of the two dashed lines).
Fig. 4. The vis (A) and FT-IR images of the water (B) and oil (C) flows in the microchannels, generated based on the water vibration band at 3380 cm−1 and the oil vibrationbands at 1143 cm−1. Scale bars represent 100 �m. MIR absorption spectrum of water and oil (D), extracted from the main and the second channel (at the cross of the twodashed lines).
Actuators B 196 (2014) 175–182 179
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E. Polshin et al. / Sensors and
enerated based on the absorption of the PDMS layer at 2962 cm−1
B) and 1260 cm−1 (C).According to the Rayleigh criterion the spatial resolution of the
btained image was increased from 9.67 �m to 4.11 �m when mov-ng from lower (1260 cm−1) to higher (2962 cm−1) wavenumbers4]. The MIR spectrum was extracted from the upper corner of theT-IR image and is displayed in Fig. 3(D). The MIR spectrum showedhe characteristic absorption bands of PDMS functional groups.wo peaks at 2962 cm−1 and 2902 cm−1 can be assigned to methyltretching vibrations, while peaks at 1446 cm−1 and 1409 cm−1 cor-espond to methyl bending vibrations [14]. The main peaks of PDMSase monomer were found at 1260 cm−1 and between 1130 cm−1
nd 1000 cm−1, and correspond to Si CH3 and Si O Si stretch-ng vibrations, respectively [15].
Second, the channels of the IR chip were filled with water and oilo demonstrate the distribution of different chemical componentsnside the microchannel based on their MIR spectra (Fig. 4).
Despite the fact that the path length in the microfluidic chip wasimited to 10 �m (height of the channels), the absorption bands inhe spectral range of O H stretching vibration of water were verytrong and this could jeopardize quantitative analysis. However, auantitative analysis in aqueous solutions is possible in the regionelow 1800 cm−1, because the O H bending vibration band did notause signal saturation. A sinusoidal pattern was observed in thepectra extracted from the chemical images (Figs. 3(D) and 4(D)).he presence of this pattern could be explained by the effect ofnterference fringes caused by the interference between the IR radi-tion that was transmitted through the chip and radiation that waseflected internally [10]. The effect of fringes in the MR spectraan reduce the accuracy of the analysis that can be achieved byhe transmission FT-IR spectroscopy. The complete elimination ofhis effect is extremely difficult, but the amplitude of the interfer-nce fringes can be reduced using wafers with a similar refractivendex to that of the sample. Nevertheless, the FT-IR images cre-ted with the FPA camera showed that it is possible to visualize theistribution of different chemical components in the microfluidicevice and to extract important spectral information relevant forualitative and quantitative analyses.
nzyme kinetic study
Glucose oxidase catalyzes the oxidation of �-d-glucose to d-lucono-1,5-lactone and hydrogen peroxide in the presence ofolecular oxygen [16,17]:
First, the MIR absorption spectra of the components involvedn the reaction between glucose and glucose oxidase were col-ected. Fig. 5 shows the spectra of the substrate (d-glucose), enzymeglucose oxidase) and the reaction product (d-glucono-1,5-lactone)
easured with ATR/FT-IR.The main absorption bands of d-glucose (A) arise from C O
nd C C stretching vibrations [11]. The MIR spectrum of GOX (B)hows the absorption of Amide I, II and III bands at 1650, 1548nd 1250 cm−1, respectively, C H bending vibrations in the region450–1350 cm−1 and C O stretching vibration at 1077 cm−1 [18].he spectrum of d-glucono-1,5-lactone has a very characteristicrequency at 1740 cm−1 which is assigned to its C O stretch-ng vibration [11], while the peak at 1230 cm−1 is caused by the
O stretching vibration [19]. Both the C O (1740 cm−1) and C O
1230 cm−1) bands are located outside the main absorption bandsf d-glucose and glucose oxidase. Therefore, these two peaks weresed as markers of product formation using the FT-IR differencepectroscopy.
Fig. 5. MIR spectra of 100 mM glucose (A), 45 �M glucose oxidase (B), and 100 mMd-glucono-1,5-lactone (C).
Second, the oxidation of d-glucose by GOX was followed in realtime with the ATR/FT-IR and the on-chip set-up as described in thesection “FT-IR measurements”. The absorption changes that weretaking place during the enzymatic reaction with GOX are shownin Fig. 6 for both the ATR accessory (A) and the on-chip configura-tion (B). The figures represent the difference in absorbance betweenthe background spectrum and the sample spectra shown for 5 minintervals. Negative absorption bands of d-glucose can be seen in thespectra obtained from both the �ATR accessory and on-chip FT-IRmicroscopy. They indicate mainly the consumption of the substrateduring the course of the reaction. As gluconolactone has absorptionbands in the spectral range between 1200 and 950 cm−1, most likelythe changes in this domain were influenced both by increasingabsorption of gluconolactone and decreasing absorption of glucose.For both ATR/FT-IR and on-chip FT-IR microscopy it can be clearlyseen how the product peaks at 1740 and 1230 cm−1 are increasingas a function of time, demonstrating the formation of the productd-glucono-1,5-lactone.
A small discrepancy in the spectral information was observedwith respect to the negative absorption pattern of d-glucosemeasured with the ATR and on-chip strategy. Presumably, this dif-ference comes from the different times at which the reaction wasinitiated. In case of ATR, the reagents were first mixed and then the
background was recorded, while in case of on-chip measurementsthe components of the reaction were mixed by diffusion in thechannel after recording the background spectrum of the flow con-taining the educts. Therefore, the background spectrum collected
180 E. Polshin et al. / Sensors and Actuators B 196 (2014) 175–182
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ig. 6. The kinetic difference MIR spectra of the enzymatic reaction between d-gluT-IR microscopy (B). The number of each spectrum corresponds to the following ti
sing ATR already comprised some bands of the product that couldhange the position and shape of the negative bands.
As can be seen from Fig. 6 the product bands at 1740 and230 cm−1 appeared earlier for the ATR approach indicating alower reaction rate for the on-chip configuration. One of the rea-ons for that is the method for mixing the components, i.e., fastonvective mixing for ATR versus diffusion based mixing for on-hip, which is known to be much slower for large biomoleculessuch as proteins) in comparison to small molecules in aqueousolutions [7]. Therefore, the rate of the d-glucose consumption wasot only determined by the kinetics of the GOX reaction itself,ut was also influenced by the diffusion of the components in theicrochannels. Another reason for slower progress of the reaction
n-chip can be related to the oxygen level, which increases the reac-ion rate when it is present at higher levels. Obviously it was lowern the channel of the microfluidic device compared to ATR crys-al, which was not closed from the environment during the kinetic
easurements.The kinetic parameters of glucose oxidase were determined
ased on the Michaelis–Menten equation, which is widely usedn enzymology for the determination of kinetic constants. The KM
alue for on-chip assay can be considered as an ‘apparent’ KM sincet also includes the effect of diffusion of the reaction reagents in the
ig. 7. The Michaelis–Menten curves for the ATR approach (A) and on-chip configuratio230 cm−1.
100 mM) and glucose oxidase (22.5 �M) as followed by ATR/FT-IR (A) and on-chipmin): 1 = 0.16; 2 = 5.0; 3 = 10.0; 4 = 15.0; and 5 = 20.0.
microchannels. The reaction was followed at constant concentra-tion of GOX (22.5 �M). The concentration of d-glucose ranged from12.5 to 300 mM for the ATR accessory and from 12.5 to 150 mMfor on-chip approach. Fig. 7 shows the Michaelis–Menten curvesfor ATR/FT-IR and on-chip FT-IR microscopy for two wavelengths(1740 and 1230 cm−1) that were characteristic for the product for-mation during the course of the reaction (appearance of C O andC O vibrations).
The progress of the reaction gradually increased due to anincreasing concentration of the substrate, until the reaction ratestabilized. For ATR, it was observed that the reaction rate wasalready stable at 150 mM of d-glucose (Fig. 7). At higher concen-tration levels of the substrate there was no further increase in thereaction rates as could be derived from both absorption peaks at1740 and 1230 cm−1. Therefore, to save time for on-chip measure-ments the reaction was followed till 150 mM of d-glucose. Theestimated Michaelis–Menten constants (KM) and the maximumrates of the reaction (Vmax) are shown in Table 1.
It was found that the maximum rate of the reaction was higher
for ATR in comparison to the on-chip FT-IR microscopy assay. Asexplained above, this was probably due to different mixing meth-ods and oxygen concentrations. A minor variation was found in thereaction rates for the two wavenumbers – 1740 and 1230 cm−1.
n (B) based on the product vibration bands (d-glucono-1,5-lactone) at 1740 and
E. Polshin et al. / Sensors and Actuators B 196 (2014) 175–182 181
Table 1Michaelis–Menten parameters of glucose oxidase for the ATR accessory and on-chip configuration.
ATR On-chip
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Wavenumber (cm ) 1740
Vmax (mM/min) 8.7 ± 0.5
KM (mM) 26.0 ± 7.4
he exact reason for this is not known, but it could be related toariation of the absorption intensities and extinction coefficients,hich are different for different wavelengths. The KM values, on
he other hand, were comparable for both ATR and on-chip mea-urements. However, a larger difference was observed betweenhe two wavenumbers for the on-chip assay. Several articles haveeported kinetic parameters of GOX; the KM and Vmax values varymong different studies. For example, the KM value of GOX from. favus was 10.9 mM [20], while Witt et al. [17] and Sukhachevat al. [21] reported a lower KM value of 5.7 mM and 3.3 mM forenicillium amagasakiense ATCC 28686 and Penicillium funiculosum33, respectively. Different experimental conditions (pH, temper-ture, buffer type, etc.) and the diffusion effects could result in thisifference of KM values. On the other hand, the Michaelis–Mentenonstant of GOX from A. niger was found to be 33 mM [22], whichs in a good agreement with the values obtained in this researchsing both ATR/FT-IR and on-chip FT-IR microscopy (Table 1). Thealues for Vmax are even more difficult to compare between dif-erent studies as they depend on the reaction conditions in eacharticular case (e.g., the temperature, the origin of the enzyme, theH, etc.). The literature reports numbers for Vmax of GOX between50 and 1000 U/mg [23]. In future research, the optimization of thenzymatic assay should be carried out with respect to size of theicrofluidic channels, mixing of the components, flow rates, pH,
emperature, buffer, etc.
onclusions
In this work, the integration of microfluidics with FT-IRicroscopy was realized. A microfluidic chip was built from IR
ransparent materials (Si and CaF2) using standard photolithogra-hy techniques. The chemical images of the continuous water andil flows in the microfluidic device were acquired with an imagingPA detector, revealing the spatially resolved distribution of chem-cal components in the microchannels. A single point detector wassed for the enzymatic assay of glucose oxidase from Aspergillusiger executed on a microfluidic chip. The obtained results showhat the reaction rate was limited by diffusion for on-chip FT-IR
icroscopy compared to the ATR approach and resulted in slowerrogress of the reaction. Nevertheless, the presented IR chip cou-led to FT-IR microscopy could be applied as a novel label-freeetection concept for various enzymatic assays, especially the oneshat cannot be directly studied by conventional UV or vis spec-roscopy.
cknowledgments
We gratefully acknowledge the financial support of the KU Leu-en Research Council for the post-doctoral mandate of EP and theDO-project CellPhinder. We thank the ESAT-MICAS research teamProf. R. Puers and Dr. F. Ceyssens) for the assistance in the micro-abrication of the chip. DDV is grateful to the Hercules Foundationproject 08/39; FTIR mapping microscope).
eferences
[1] A. Ríos, A. Escarpa, M.C. González, A.G. Crevillén, Challenges of analyticalmicrosystems, TrAC: Trends in Analytical Chemistry 25 (2006) 467–479.
[2] D. Mark, S. Haeberle, G. Roth, F. von Stetten, R. Zengerle, Microfluidic lab-on-a-chip platforms: requirements, characteristics and applications, ChemicalSociety Reviews 39 (2010) 1153–1182.
[3] K.L.A. Chan, S. Gulati, J.B. Edel, A.J. de Mello, S.G. Kazarian, Chemical imag-ing of microfluidic flows using ATR-FTIR spectroscopy, Lab on a Chip 9 (2009)2909–2913.
[5] K.L.A. Chan, S.G. Kazarian, FT-IR spectroscopic imaging of reactions inmultiphase flow in microfluidic channels, Analytical Chemistry 84 (2012)4052–4056.
[6] M. Kakuta, P. Hinsmann, A. Manz, B. Lendl, Time-resolved Fourier transforminfrared spectrometry using a microfabricated continuous flow mixer: appli-cation to protein conformation study using the example of ubiquitin, Lab on aChip 3 (2003) 82–85.
[7] W. Buchegger, C. Wagner, B. Lendl, M. Kraft, M.J. Vellekoop, A highly uniformlamination micromixer with wedge shaped inlet channels for time resolvedinfrared spectroscopy, Microfluidics and Nanofluidics 10 (2010) 889–897.
[8] W. Buchegger, C. Wagner, P. Svasek, B. Lendl, M. Kraft, M.J. Vellekoop, Fab-rication and characterization of a vertical lamination micromixer for mid-IRspectroscopy, Sensors and Actuators B: Chemical 159 (2011) 336–341.
[9] C. Wagner, W. Buchegger, M. Vellekoop, M. Kraft, B. Lendl, Time-resolvedmid-IR spectroscopy of (bio)chemical reactions in solution utilizing a newgeneration of continuous-flow micro-mixers, Analytical and BioanalyticalChemistry 400 (2011) 2487–2497.
10] P.R. Griffiths, J.A. de Haseth, Fourier Transform Infrared Spectrometry, 2nd ed.,John Wiley & Sons Ltd., Hoboken, NJ, USA, 2007.
11] K. Karmali, Karmali A., A. Teixeira, M.J. Curto, Assay for glucose oxidase fromAspergillus niger and Penicillium amagasakiense by Fourier transform infraredspectroscopy, Analytical Biochemistry 333 (2004) 320–327.
12] S. Kumar, A. Barth, Following enzyme activity with infrared spectroscopy, Sen-sors (Basel, Switzerland) 10 (2010) 2626–2637.
13] J. Voldman, M.L. Gray, M.A. Schmidt, Microfabrication in biology and medicine,Annual Review of Biomedical Engineering 1 (1999) 401–425.
14] S.A. Merschman, S.H. Lubbad, D.C. Tilotta, Poly(dimethylsiloxane) films as sor-bents for solid-phase microextraction coupled with infrared spectroscopy,Journal of Chromatography A 829 (1998) 377–384.
15] P. Jothimuthu, A. Carroll, A.A.S. Bhagat, G. Lin, J.E. Mark, I. Papautsky, Pho-todefinable PDMS thin films for microfabrication applications, Journal ofMicromechanics and Microengineering 19 (2009) 045024.
16] R. Wilson, A.P.F. Turner, Review article glucose oxidase: an ideal enzyme,Biosensors & Bioelectronics 7 (1992) 165–185.
17] S. Witt, G. Wohlfahrt, D. Schomburg, H.J. Hecht, H.M. Kalisz, Conserved arginine-516 of Penicillium amagasakiense glucose oxidase is essential for the efficientbinding of beta-d-glucose, Biochemical Journal 347 (2000) 553–559.
18] M. Portaccio, B. Della Ventura, D.G. Mita, N. Manolova, O. Stoilova, I. Rashkov,M. Lepore, FT-IR microscopy characterization of sol–gel layers prior and afterglucose oxidase immobilization for biosensing applications, Journal of Sol–GelScience and Technology 57 (2010) 204–211.
19] J. Coates, Interpretation of infrared spectra: a practical approach, in: R.A. Meyers(Ed.), Encyclopedia of Analytical Chemistry, John Wiley& Sons Ltd., Chichester,UK, 2000, pp. 10815–10837.
20] K.K. Kim, D.R. Fravel, G.C. Papavizas, Production purification, and propertiesof glucose-oxidase from the biocontrol fungus Talaromyces-flavus, CanadianJournal of Microbiology 36 (1990) 199–205.
21] M.V. Sukhacheva, M.E. Davydova, A.I. Netrusov, Production of Penicilliumfuniculosum 433 glucose oxidase and its properties, Applied Biochemistry andMicrobiology 40 (1) (2004) 25–29.
22] B.E.P. Swoboda, V. Massey, Purification properties of glucose oxidase fromAspergillus niger, Journal of Biological Chemistry 240 (5) (1965) 2209–2215.
Evgeny Polshin was born in Russia, in 1983. He obtained his bachelor (2004) andmaster (2006) degree in chemistry at Saint-Petersburg University. In 2007 he was
an international scholar at University of Leuven (Belgium) in the framework of abilateral scientific collaboration between Russia and Belgium. He received the PhDdegree in bioscience engineering from the University of Leuven in 2012. His scien-tific interests include FT-IR spectroscopy, microscopy and imaging, continuous anddigital microfluidics.
the biosensor group. His main research interests involve the development of novel
82 E. Polshin et al. / Sensors and
ert Verbruggen was born in Belgium, in 1985. He obtained his bachelor (2006) andaster (2008) degree in bio-engineering sciences, with specialization in environ-ental technologies and bio-nano technologies, at the University of Leuven. In 2008
e started his PhD research at the biosensor group of the MeBioS department at theniversity of Leuven. His key subjects are digital microfluidics including modeling,agnetic microparticles, microscopy and imaging.
aan Witters was born in Herk-de-Stad, Belgium, in 1986. He received his BS and MSn bio-engineering from the KU Leuven, Leuven, Belgium in 2009. His master thesis
as in the field of enzymatic assays performed on microfluidic systems. He obtainedis PhD at KU Leuven in October 2013 and his research interests concern the designnd application of digital microfluidic systems for performing high-throughput bio-ssays and miniaturized materials synthesis.
ert Sels obtained his PhD degree in 2000 at the Catholic University of Leuven onxidation chemistry, after which he did a post-doc with BASF until 2002. Another
years post-doc for the National Science Foundation was dedicated to the “activa-ion of nitrous oxide” and the “microscopic imaging of catalytic events”. He becamessistant professor in 2003, teaching courses on analytical chemistry and heteroge-eous catalysis. He is appointed full professor at Leuven since 2006 in the Facultyf Bioengineering Sciences. He has published about 140 scientific papers and 11atents, and is recipient of numerous awards including the prestigious internationalSM chemistry award. His current research explores heterogeneous catalysis for
enewables conversion and small molecule activation.
irk De Vos is a full professor in bioscience engineering at KU Leuven. His mainnterests lie in porous materials, the interactions of organic molecules with these
aterials and in catalytic transformations inside the pores. Together with J. Hofkens
tors B 196 (2014) 175–182
and M. Roeffaers (both from KU Leuven), he pioneered the visualization of activesites in porous materials using light microscopic techniques. His team made ground-breaking discoveries in the application of metal-organic frameworks to liquid phaseseparation and catalysis. His awards include the BASF Catalysis Award and the D.W.Breck Award of the International Zeolite association.
Bart Nicolai obtained a MSc in bioscience engineering from Gent University(Belgium) in 1986, and a MSc in applied mathematics (1988) and PhD in bioscienceengineering (1994) from the University of Leuven (Belgium). He is now full professorat the latter institute. Since 2005 he is head of the division mechatronics, biostatis-tics and sensors (MeBioS) of the biosystems department at the University of Leuven.In addition, he is responsible for co-ordinating the research in the Flanders Center ofPostharvest Technology (VCBT), an experimental facility which was established as apublic–private partnership between the K University of Leuven and the Associationof Belgian Horticultural Auctions in 1997. His main research interests are posthar-vest biology and technology, heat and mass transfer, and fruit and vegetable qualitywith flavor in particular.
Jeroen Lammertyn is MSc in applied biological sciences and MSc in biostatistics.He obtained his PhD in applied biological engineering at University of Leuven in2001. In 2002–2003 he was research associate at the Pennsylvania State University,USA. Since October 2005 he is professor at the University of Leuven and head of
bio-molecular detection concepts and miniaturized analysis systems: bio-assaydevelopment (e.g., aptamers, biofunctionalized nanomaterials), optical sensors (e.g.,fiber optic SPR sensors, FT-IR and NIR spectroscopy), micro- and nanofluidics (e.g.,lab-on-a-chip technology).
