Immobilization of multi-enzyme microreactors inside microfluidic devices

Sensors and Actuators B 106 (2005) 335–342

Immobilization of multi-enzyme microreactors inside microfluidic devices Won-Gun Koha , Michael Pishkoa,b,c,∗ a

Department of Chemical Engineering, 104 Fenske Laboratory, The Pennsylvania State University, University Park, PA 16802-4420, US b Department of Chemistry, The Pennsylvania State University, University Park, PA 16802-4420, US c Department of Materials Science & Engineering, The Pennsylvania State University, University Park, PA 16802-4420, US Received 23 March 2004; received in revised form 6 August 2004; accepted 13 August 2004 Available online 28 September 2004

Abstract A simple method to fabricate enzyme-containing microscopic hydrogel structures in microfluidic devices for the potential use in micro total analysis systems (␮-TAS) is described. Poly(ethylene glycol)-based hydrogel microstructures were prepared inside microchannels by photolithography and enzymes conjugated to a pH sensitive fluorophore (SNAFL-1) were incorporated into these hydrogel microstructures. Because of the ratiometric pH-dependent nature of SNAFL fluorescence, hydrogel microstructures exhibited a different emission intensity ratio with pH and this intensity ratio changed almost linearly between pH 7 and 12. When alkaline phosphatase-containing microreactors were exposed to p-nitrophenylphosphate (pNPP) as a substrate, phosphoric acid was produced inside the microstructure by enzymatic-catalyzed hydrolysis of the substrate and subsequently decreased the microenvironment pH. Because of the relatively rapid mass transport of analyte through the hydrogel, enzyme-catalyzed reaction was easily detected by change in emission intensity ratio before and after exposure to substrates. Enzyme-catalyzed reactions were quite fast and reached 90% of maximum value within 10 min. Data were analyzed using a modified Michaelis–Menten equation and apparent Michaelis constants could be obtained. This system was also successfully applied to urea hydrolysis by urease. © 2004 Elsevier B.V. All rights reserved. Keywords: ␮-TAS; Hydrogel; Microfluidic device; Enzyme reaction; Fluorescence

1. Introduction Miniaturization of analytic devices in micro total analysis systems (␮-TAS) represents a natural extension of microfabrication technology to chemistry and biology with applications in high throughput screening and in portable analytical measurement devices. The recent developments in ␮-TAS have allowed the miniaturization and integration of (bio)chemical instruments into a microchip-like format (“labon-a-chip”). The miniaturization of chemical reactors to the micrometer scale creates various advantages over benchtop instruments, including smaller dead volume and sample consumption, shorter analysis time, low cost, greater sensitiv∗

Corresponding author. Tel.: +1 814 863 4810; fax: +1 814 865 7846. E-mail address: [email protected] (M. Pishko).

0925-4005/$ – see front matter © 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.snb.2004.08.025

ity and the capability performing on simultaneous reactions [1–6]. These microanalysis devices can be classified into two categories based on the complexity of the fluidics involved. One category is microarray-based microdevices where substances such as DNA and protein are immobilized on the chip. The second category is microfluidics-based microdevices where substances are transported, reacted and separated on chip. Microfluidic devices offer potential analytical advantages over planar array microchip such as enhanced mass and heat transfer, lower sample volumes, and ease of integration with miniaturized sample preparation modules [7,8]. Several investigators have explored the role that enzymes could play in lab-on-a-chip technologies and have investigated enzyme-substrate reactions within microfabricated channel networks using electrokinetic or pressure-driven flow

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[9–12]. A conventional type of microfluidic device for enzyme assay consists of several reagent wells each connected by channels to a main channel where the reaction and analysis regions are located [13–15]. Recently, however, a few groups have tried to immobilize enzymes inside microchannels for assay development [16–18]. Ismagilov et al. also investigated enzyme reaction using fluid–fluid diffusional contacts using microfluidic arrays [19]. Hydrogels have been used in biology and medicine for many years because of their properties of hydrophilicity, biocompatibility, permeability and mechanical strength [20]. Highly cross-linked hydrogel networks are capable of encapsulating protein or mammalian cells and have been used for numerous applications such as biosensors, cell encapsulation and drug delivery [21–25]. Previously, we developed PEG hydrogel microbead or microarray containing enzymes for use as optical sensors and could detect up to micromolar concentration of analytes [26,27]. PEG hydrogels provided a protective environment for the immobilized enzyme and inhibited degradation and fouling. Since Beebe et al. proposed fabrication of hydrogel microstructures inside microchannels for the use as microactuators [28], several groups have tried to make hydrogels containing DNA or mammalian cells inside microchannels for the DNA hybridization or potential drug screening systems [28–31]. In this study, we fabricated enzyme-containing poly(ethylene glycol) (PEG) hydrogel microstructures in microfluidic channels and performed enzyme assays using carboxy seminaphthofluorescein (SNAFL-1) conjugated to alkaline phosphatase and urease in PEG hydrogel microstructures for the potential use of this systems as biosensors or microreactors.

2. Materials and methods

sodium phosphate dibasic heptahydrate, and 0.15 M NaCl in 18 M cm deionized water (Milli-Q Ultrapure, Millipore) 2.2. Preparation of SNAFL labeled enzyme solution Using an established protocol, SNAFL-enzyme conjugates were prepared by reacting 1 mg SNAFL-1 succinimidyl ester dissolved in 100 ␮L of DMSO with the enzyme dissolved in a 100 mM PBS solution (pH 8.2). Unreacted dye was separated by overnight dialysis [32]. The final enzyme concentration was approximately 2 mg/mL. 2.3. Fabrication of microfluidic device Microchannels in PDMS were obtained by curing a 10:1 mixture of PDMS prepolymer and curing agent against a Si master which has a negative pattern of the desired microchannel defined with SU-850 negative photoresist (Microlithography Chemical Corp, Newton, MA). After cured for several hours at 60 ◦ C, PDMS replica was removed from the master and oxidized in an oxygen plasma (Harrick Scientific Co., Ossining, NY) with glass slide for 1 min. Bringing the oxidized PDMS and glass slide into conformal contact resulted in an irreversible seal and thus formed an enclosed microchannel. These microchannels were treated with dilute TPM solution in perfluorooctane for 10 min immediately after sealing to enhance the adhesion of hydrogel microstructure inside the microchannels. To make inlet and outlet ports in the microfluidic device, several holes were punched through PDMS replica using 16-gauge needle. Polyethylene tubes were inserted into these holes and then connected to syringe pump (Harvard Apparatus, Holliston, MA) to complete the microfluidic device. These microfluidic devices were mounted on the stage of a microscope for real-time fluorescence detection and imaging. This experimental apparatus is illustrated in Fig. 1.

2.1. Chemicals and materials Alkaline phosphatase (AP) from E. coli, urease, pnitrophenylphosphate (pNPP) and urea as substrates for the enzymes, fluorescein isothiocyanate (FITC) and tetramethylrhodamine isothiocynate (TRITC) were purchased from Sigma Chemical Co. (St. Louis, MO). Poly(ethylene glycol) diacrylate (PEG-DA, MW 575) and perfluorooctane was obtained from Aldrich Chemical Co. (Milwaukee, WI). 2Hydroxy-2-methyl-1-phenyl-1-propanone (Ciba, Tarrytown, NY) was used as a photoinitiator. 5-(and-6)-carboxy SNAFL1 was purchased from Molecular Probes (Eugene, OR). Poly(dimethyl siloxane) (PDMS) elastomer was purchased as Dow Corning Sylgard 184 (Midland, MI), which is composed of a prepolymer and curing agent. 3-(Trichlorosilyl)propyl methacrylate (TPM) was purchased from Fluka Chemicals (Milwaukee, WI). The photomasks for making hydrogel patterns were purchased from Advanced Reproductions (Andover, MA). Phosphate buffered saline (PBS, 0.1 M pH 7.4) consisted of 1.1 mM potassium phosphate monobasic, 3 mM

2.4. Fabrication of PEG hydrogel microstructures inside microchannels PEG hydrogel microstructures were fabricated from PEGDA (MW 575) as the base macromer. The gel precursor solution was prepared by dissolving 10 ␮L of photoinitiator per 1 mL of PEG-DA solution. When fluorescent hydrogel structures were to be prepared, 50 ␮L of SNAFL or SNAFLlabeled enzyme solution per milliliter of precursor solution was also added. The microchannels were filled with these precursor solutions and then exposed to 365 nm, 300 mW/cm2 UV light (EFOS Ultracure 100ss Plus, UV spot lamp, Mississauga, Ontario) for 1 s through a photomask on the top of glass slide. After UV photopolymerization, only exposed polymer regions underwent free radical cross-linking and became insoluble in common PEG solvents such as water. Finally, by flushing the channel with PBS, desired hydrogel structures were obtained inside a microchannel. The final enzyme concentration within the gel was approximately 0.1 mg/mL.

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Fig. 1. Experimental apparatus for the detection of enzyme-catalyzed reactions in PEG hydrogel microstructures.

2.5. Image acquisition and analysis A Zeiss Axiovert 200 microscope equipped with an integrated color CCD camera (Carl Zeiss Inc., Thornwood, NY) was used to obtain optical and fluorescence images of hydrogels inside microchannels. For fluorescence microscopy, fluorescein isothiocyanate (FITC) and tetramethylrhodamine isothiocynate (TRITC) filters with excitation and emission wavelengths of 480 ± 40 nm/535 ± 50 nm (FITC) and 545 ± 30 nm/620 ± 60 nm (TRITC) were used. Fluorescence intensity was quantified as a function of substrate concentration using the ratio of intensity of emission at 535 nm compared to 620 nm, representing the acidic and basic forms of SNAFL-1. All data analysis was performed using commercially available image analysis software (Zeiss Image or KS 300, Carl Zeiss Inc., Thornwood, NY)

3. Results and discussions 3.1. PEG hydrogel microstructures inside microchannels Hydrogel microstructures were fabricated inside microchannels using photolithography. Using a wellestablished method to create microfluidic devices [33], an approximately 100–300 ␮m wide, 50 ␮m deep microchannels were created in PDMS and irreversible sealing to a glass substrate enabled fluid to be pumped through microchannels without leakage. The formation of PEG hydrogels is based on the UV initiated free-radical cross-linking of acrylate end groups on PEG-DA, resulting in the formation of polyacry-

late networks highly cross-linked with PEG [21]. To avoid deformed structures, the introduced precursor solution was allowed to reach a stationary state and then exposed to UV light. Various shapes and sizes of hydrogel microstructures could be aligned with microchannels using different masks. Fig. 2 shows the hydrogel microstructures containing FITC inside a microchannel. As shown in Fig. 2(a), only UV-illuminated regions underwent photopolymerization and gelled inside the microchannel, and the fluorescence image shows that unreacted precursor solution was totally removed by flushing channel with water or PBS. (Fig. 2(b)) After hydrogel structures were fabricated inside microchannels, all fluids were pumped into microchannels at flow rates ranging from 5 to 20 ␮L/min by syringe pump and at these flow rates, hydrogel microstructures remained in place without buckling and migrating in the channels. Longer UV exposure time resulted in hydrogel structure larger than the feature size of the mask, because of the mass transfer of free radical species outside the illuminated area and their subsequent reaction with PEG-DA [27]. In this experiment, the polymerization process took only 1 s and this short exposure time allowed us to obtain hydrogel microstructures as small as 7 ␮m. We also fabricated hydrogel microstructures containing different compositions in a single channel. First, microstructures containing TRITC were made inside the channel (Fig. 3(a)) and then precursor solution containing FITC was polymerized next to TRITC-containing hydrogels as shown in Fig. 3(b). In these images, we observed that a small amount of FITC diffused into TRITC-containing hydrogels from the weak green light from hydrogel microstructures on

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Fig. 2. Images of PEG hydrogel microstructures containing FITC inside microchannels: (a) transmission microscopy; (b) fluorescence microscopy.

the left side. Finally Fig. 3(c) shows the combined image of (a) and (b). Based on this result, we anticipated we can fabricate hydrogel microstructures containing different types of enzymes in single channel for multianalyte biosensing. 3.2. pH response of SNAFL in PEG hydrogel SNAFL, a ratiometric pH-dependent fluorophore, has both dual emission and dual excitation properties from both the acid and base forms. The pKa of SNAFL is approximately 7.8. In the acidic condition, free dye is optimally excited at 510 nm, with an emission maximum at 545 nm and in a basic environment, the optimum excitation and emission values are red shifted to 542/645 nm. Therefore both intensity and emission wavelengths change with changes in pH and this dualwavelength feature of SNAFL make it particularly suitable for ratiometric sensing, potentially reducing error associated with photobleaching and lamp intensity fluctuations. To investigate the pH response of SNAFL in PEG hydrogel microstructures, a cylindrical array of hydrogel microstruc-

Fig. 3. Microchannels containing four hydrogel microstructures (two labeled with FITC and two labeled with TRITC): (a) fluorescence images obtained by TRITC filtered light; (b) FITC filtered light; and (c) composite images.

tures containing SNAFL was fabricated inside microchannels and buffer solutions of different pH were introduced to each microchannel. Fig. 4 shows two hydrogel-containing microchannels exposed to different pH solutions. Here, upper channel was filled with pH 11 solution, while lower channel was filled with pH 7 solution. As shown in this figure, hydrogel microstructures in pH 7 solution showed slightly acidic spectrum, with strong emission through the FITC filter and weak emission through the TRITC filter, resulting in a yellow-green color when two images were overlapped. However, after exposure to a buffered solution at pH 11, hydrogel fluorescence underwent a strong red shift and the overlapped image appeared orange. Photobleaching of the dye was not observed during UV exposure or during repeated excitation over 1 week period. For quantitative analysis, changes in flu-

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Fig. 5. pH response of SNAFL in PEG hydrogel microstructures. Fluorescence intensity was quantified as the ratio of the emission at 535 nm compared to 620 nm.

variation in intensity between identical microstructures on a given substrate was less than 5%. 3.3. Characterization of reactions inside PEG microstructures in microfluidic devices

Fig. 4. Hydrogel-containing microchannels exposed to different pH solutions (pH 7 for upper channel and pH 11 for low channel): (a) fluorescence image obtained with FITC filter; (b) TRITC filter; and (c) composite image of obtained with FITC and TRITC filters.

orescence emission intensity of SNAFL-containing PEG hydrogel microstructures were measured as a function of pH. Here, fluorescence intensity was quantified as the ratio of the emission at 535 nm compared to 620 nm representing the acidic and basic forms of dye. As shown in Fig. 5, the intensity ratio changed with pH almost linearly between pH 7 and 12, and at this pH range, intensity ratio decreases with pH because of the emission red shift in a basic environment. The

To investigate the potential use of these microstructures in a micro-total-analysis-system (␮-TAS) or in microreactors for synthesis, SNAFL-conjugated alkaline phosphatase was incorporated in hydrogel microstructures inside microfluidic channels and pNPP as a substrate was introduced to microchannels using a syringe pump. Before the substrate was introduced to channels, hydrogel microstructures inside channel were exposed to substrate-free PBS solution until emission intensity ratio reached steady state. Even though the optimum pH for alkaline phosphatase activity is approximately 8, more basic buffer solution (pH 10) was used because of an acidic shift in the reported SNAFL pKa due to the PEG hydrogel microenvironment pH [26]. We found that there was a different in fluorescence emission intensity at both emissions peaks between hydrogels before and after enzymecatalyzed reaction. That is, hydrogel microstructures exposed to substrate-containing buffer had stronger green emission and weaker red emission than those prior to substrate introduction. This results from the decrease of microenvironment pH in the gel through the production of phosphoric acid as a result of the enzyme-catalyzed reaction, indicating that the enzyme retained its activity after the UV photopolymerization process. Fig. 6(a) shows the change of intensity ratio (535 nm/620 nm) as a function of substrate concentration. As expected, the change in emission intensity ratio increased with substrate concentration because microenvironment pH in hydrogel microstructure decreased by the production of phosphoric acids. The change in emission intensity ratio reached maximum value at high substrate concentration. In

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Fig. 6. Change of fluorescence emission intensity ratio by enzyme-catalyzed reaction: (a) plot of normalized fluorescence intensity ratio against concentration of pNPP; (b) Lineweaver–Burk plot for the reaction of alkaline phosphatase (R2 = 0.96); (c) plot of normalized fluorescence intensity ratio against concentration of urea; and (d) Lineweaver–Burk plot for the reaction of urease (R2 = 0.95).

an earlier study, we demonstrated that mass transfer is rapid enough for sensing application in photopolymerized micronsized PEG hydrogel that contain analyte sensitive recognition molecules [34]. Besides alkaline phosphatase, other enzyme-catalyzed reaction was also investigated using these microfluidic devices. Fig. 6(c) shows the reaction of immobilized urease upon exposure to the urea. As shown here, the urease-catalyzed reaction caused a change in pH toward basic range due to the formation of NH4 + . These results from two enzymes demonstrated the ability of this system to sense in both acidic and basic ranges of pH. Experiments conducted showed the sensor response time was quite rapid that change of intensity ratio by enzymecatalyzed reaction reached 90% of maximum value within 10 min. The lowest concentration of substrates analyzed experimentally in this study is 10 ␮M. However, much lower concentration of substrates could be detected in this system considering the detection limit of 16 nM in previous SNAFLenzyme biosensor [26].

Simple enzyme kinetics have been commonly evaluated using the Michaelis–Menten equation, V0 =

Vmax [S] Km + [S]

This equation relates the initial rate of the enzyme reaction (V0 ) to the concentration of substrate ([S]), a maximum rate (Vmax ) and the Michaelis constant (Km ). The values of Vmax and Km were obtained from a reciprocal Lineweaver–Burk plot. In this study, the steady state reaction rate as a function of substrate concentration was expressed as the normalized steady state intensity ratio (I0 ). Therefore, a modified Michaelis–Menten equation can be expressed as following equation, I0 =

Imax [S] Km(app) + [S]

where Imax is the maximum value of normalized intensity ratio and Km(app) is the apparent Michaelis constant. The ki-

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netic constants Km measured with immobilized enzymes are not true kinetic constants equivalent to those obtained in free solution. It is an apparent Km because the diffusion of substrates from the bulk solution to the microenvironment of an immobilized enzyme limits the rate of the enzyme reactions. Therefore, apparent Michaelis constant was used here. A plot of the Lineweaver–Burk equation then yields Km(app) and Imax of this system (Fig. 6(b) and (d)), where the slope of the line is Km(app) /Imax and the intercept is 1/Imax . The results of this treatment yield Km(app) = 0.57 and 0.35 mM for alkaline phosphatase and urease, respectively. In the literature, it was reported that Km value for urease is 10.5 mM at pH 7.0, 25 ◦ C and for alkaline phosphatase is 3 mM at pH 8.2, 37 ◦ C, both in aqueous buffer rather than in a hydrogel [35]. Since the enzymes were encapsulated in cross-linked hydrogel which can hinder the mass transfer of substrates, the characterization of mass transport inside hydrogel is very important. Mass transfer in hydrogels is strongly influenced by the cross-linking density of the hydrogel, which may be characterized by calculating the number average molecular weight between crosslinks (Mc ) and mesh size. According to an earlier study, the values of Mc and mesh size for a hy˚ drogel made from PEG 575 were about 50 g/mol and 10 A respectively, and the diffusivity of small molecules such as glucose through this hydrogel were about 4.0 × 10−7 cm2 /s [21,34]. Because of the relatively small mesh size of the PEG hydrogel, no dye-labeled protein diffusion out of the gel was observed during these experiments. In a previous study, hydrogel elements based on PEG-DA (MW 575) showed about a 57% increase in volume and 50% equilibrium water content upon equilibrium in water [21]. Extensive swelling was not desirable here because swollen hydrogel microstructure may block the channel and hence fluid flow. To avoid waterinduced swelling, we added water to the precursor solution so that the resultant hydrogel had approximately its equilibrium water content and thus would not physically swell appreciably with additional water. Increased water content in precursor solution also improved diffusion of small molecular weight analytes into the hydrogel [34].

4. Conclusion Microscopic hydrogel microstructures were fabricated in a microfluidic device using photolithography for the potential use in a micro-total-analysis-systems (␮-TAS) as biosensors or microreactors. Here, we described the use of these devices to perform the continuous monitoring of enzymecatalyzed reaction using alkaline phosphatase-catalyzed hydrolysis of pNPP as a model system. Because of relatively rapid mass transport of analytes through the hydrogel, the enzyme-catalyzed reaction was easily detected by the change of emission intensity ratio before and after reaction up to micro molar concentration of substrate within several minutes.

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Other enzyme-catalyzed reaction with urease was also successfully detected using this system, demonstrating that the change in the microenvironment pH can be detected towards both the acidic and basic ranges.

Acknowledgement The work described in this paper was supported by a grant from the National Aeronautics and Space Administration (NASA, NAG 91277).

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