Selective Ion Extraction: a Separation Method for Microfluidic Devices

While electrokinetic material transport systems provide numerous benefits in the microscale movement, mixing and aliquoting of fluids, pressure-driven...
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Anal. Chem. 2002, 74, 5175-5183

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Selective Ion Extraction: a Separation Method for Microfluidic Devices Matthew B. Kerby, Michael Spaid, Spencer Wu, J. Wallace Parce, and Ring-Ling Chien*

Caliper Technologies Corporation, 605 Fairchild Drive, Mountain View, California 94043

A separation concept, selective ion extraction (SIE), is proposed on the basis of the combination of hydrodynamic and electrokinetic flow controls in microfluidic devices. Using a control system with multiple pressure and voltage sources, the hydrodynamic flow and electric field in any section of the microfluidic network can be set to desired values. Mixtures of compounds sent into a T-junction on a chip can be completely separated into different channels on the basis of their electrophoretic mobilities. A simple velocity balance model proved useful for predicting the voltage and pressure settings needed for separation. SIE provides a highly efficient separation with minimal additional dispersion. It is an ideal technique for highthroughput screening systems and demonstrates the power of lab-on-a-chip systems. Miniaturization of chemical analysis systems and the “lab-on-a chip” concept has generated a great deal of interest in the scientific community in the past decade.1-5 Many chemical and biochemical applications were developed and demonstrated using this new technology.6-8 One area of interest and commercial application is high-throughput screening (HTS) using microfluidic devices.9,10 Miniaturization reduces the amount of biological and chemical reagents used per assay and provides high quality data with high throughput. The ability to program and control fluid transport has always been the most promising feature for lab-on-a-chip devices. Electrokinetic forces have the advantages of direct control, fast response, simplicity, and allowing analytes to be selectively moved through a complex network of channels, which permits the * Corresponding author: [email protected]. (1) Manz, A. Trends Anal. Chem. 1991, 10, 144. (2) Manz, A.; Graber, N.; Widmer, H. M. Sens. Acutators 1988, 14, 101. (3) Harrison, D. J.; Manz, A.; Fan, Z.; Ludi, H.; Widmer, H. M. Anal. Chem. 1992, 64, 1926-1932. (4) Woolley, A. T.; Matthies, R. A. Anal. Chem. 1995, 67, 3676-3680. (5) Jacobson, S. C.; Ramsey, J. M. Handbook of Capillary Electrophoresis, 2nd. ed.; Landers, J. P., Ed.; John Wiley & Sons: New York, 1997; pp 827-839. (6) Micro Total Analysis System ’98; Harrison, D. J., van den Berg, A., Eds.; Kluwer Academic Publishers: Dordrecht, The Netherlands, 1998. (7) Hadd, A. G.; Raymond, D. E.; Halliwell, J. W.; Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1997, 69, 3407-3412. (8) Chiem, N.; Harrison, J. D. Anal. Chem. 1997, 69, 373-378. (9) Bousse, L.; Cohen, C. B.; Nikiforov, T.; Chow, A.; Kopf-Sill, A.; Dubrow, R.; Parce, J. W. Annu. Rev. Biophys. Biomol. Struct. 2000, 29, 155-181. (10) Sundberg, S. A.; Chow, A.; Nikiforov, T.; Wada, H. G. Drug Discovery Today 2000, 1 (HTS supplement), S42-S53. 10.1021/ac0258103 CCC: $22.00 Published on Web 09/10/2002

© 2002 American Chemical Society

implementation of a wide variety of chemical and biochemical analyses. While electrokinetic material transport systems provide numerous benefits in the microscale movement, mixing and aliquoting of fluids, pressure-driven flow avoids detrimental effects of electric fields, such as biased sampling and disruption of enzyme reactions. We previously described a universal multiport system capable of controlling pressures and voltage on multiple wells in a lab-on-a-chip microfluidic device.11 Precise flow control from each individual channel can be achieved, assuming that the hydrodynamic resistance of the network is known. In this report, we describe a separation technique that uses the combination of hydrodynamic and electrokinetic flow control to perform enzymatic assays for high-throughput screening. There are several different approaches toward assay miniaturization. One approach is based on the concept of a continuous flow assay.10 Small plugs of the preincubated substrate, enzyme, and inhibitor solution from a library of compounds are sipped onto the chip through the capillary from the microtiter plate. Buffer is sipped between each sample as a spacer. To prevent sample biasing caused by electrokinetic injection, pressure-driven flow is commonly used to transport these plugs through a network of interconnecting channels to a waste well, usually located at the end of the fluidic network, where a negative pressure (vacuum) is applied. When all of the flow on a chip is driven by a single pressure source, the hydrodynamic flow distribution or the dilution ratio in the channel network is predetermined by the fixed hydrodynamic resistances of the channels. For some assays, an electric field is also applied in part of the channel to provide separation on the basis of differences in the electrophoretic mobility of the substrate and product. An electrophoretic mobility difference between the substrate and product molecules generates a finite difference of velocity in the separation channel. This velocity change is recorded as a fluorescent intensity peak shift at the detector. Nevertheless, both substrate and product flow downstream together to the waste well. Since all reagents flow into a single waste well, the detector is usually located near the end of the separation channel in which the electric field is applied to maximize the separation power. Unfortunately, Taylor-Aris dispersion often offsets gains in peak resolution produced by simply increasing the length of the separation channel.12-14 (11) Chien, R.-L.; Parce, J. W. Fresenius’ J. Anal. Chem. 2001, 371, 106-111. (12) Taylor, G. I. Proc. R. Soc. (London) 1953, 219A, 186-203. (13) Aries, R. Proc. R. Soc. (London) 1956, 235A, 67-77.

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Several authors have investigated methods for differentially transporting ions in microfluidic devices using the combination of pressure-driven flow and electrokinetic flow. Culbertson et al. demonstrated electroosmotically induced hydraulic pumping using voltage control in a microchip with a T-junction geometry.15 Using polymer coatings to selectively reduce the electroosmotic flow in a grounded side channel, pressure-driven flows were generated via hydraulic pumping in a field-free channel. Anions were rejected from the grounded channel of the tee chip when the electrophoretic velocity of the anion out of the ground channel exceeded the pressure-driven flow into the ground channel. Cationic and neutral species passed into both channels. The ratio of flow resistances between the ground and field-free channels determined the velocity threshold for anion separation. To separate ions of different electrophoretic mobility, a separate chip was designed in which the channel dimensions were modified. In another report, Attiya et al. used a low-resistance channel in which flow rates up to 1 mL/min did not perturb a shallow channel used for electrophoretic separations with 106 times greater resistance.16 The sample introduction channel, 300 µm deep, 1 mm wide, and up to 20 mm long, had a holdup volume of 6 µL. Chen et al. has extended the same idea and proposed an electrophoretic microchip design that allows a direct coupling with hydrodynamic flowthrough analyzers for uninterrupted sampling.17 We report on a method for separating charged compounds in microfluidic devices on the basis of managing flow using multiple pressure and voltage sources. Simultaneous, independent control of the pressure-driven and electrophoretic contributions to the total velocity is possible with this approach, which determines the direction of flow of the compounds. Using multiport control, the flow velocity in any section of the microfluidic network can be set to a desired value. We present an example in which the substrate and product of an enzymatic reaction are completely separated and diverted to different wells via separate channels. This kind of combination flow control demonstrates the power of a true lab-on-a-chip system. MODEL SYSTEM To obtain a fundamental understanding of the combined pressure and electrokinetically driven species transport involved in SIE, we have used computational fluid dynamics (CFD) to simulate the analyte injection for a model T-junction geometry.18 The model geometry is three-dimensional, consisting of three 500µm-long channels labeled L1, L2, and L3 that form the branches of the T-junction, as shown in Figure 1. The cross section of the channels is rectangular, with a width of 74 µm and a depth of 12 µm. The three rectangular faces at the end of each channel segment are labeled F1, F2, and F3, as shown in Figure 1, and a subscript notation is used to reference specified quantities at each face, for example, V1 and P1 for the voltage and pressure applied across face F1. (14) Dutta, D.; Leighton, D. T., Jr. Anal. Chem. 2001, 73, 504-513. (15) Culbertson, C. T.; Ramsey, R. S.; Ramsey, J. M. Anal. Chem. 2000, 72, 2285-2291. (16) Attiya, S.; Jemere, A. B.; Tang, T.; Fitzpatrick, G.; Seiler, K.; Chiem, N.; Harrison, D. J. Electrophoresis 2001, 22, 318-327. (17) Lin, Y. H.; Lee, G. B.; Li, C. W.; Huang, G. R.; Chen, S. H. J. Chromatogr., A. 2001, 937, 115-125. (18) Bianchi, F.; Ferrigno, R.; Girault, H. H. Anal. Chem. 2000, 72, 1987-1993.

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Figure 1. Isometric view of the T-junction geometry used for the CFD calculations. The three rectangular faces at the end of each channel segment are labeled F1, F2, and F3. Each channel segment, L1, L2, and L3, is 500 µm long, with a 74-µm-wide by 12-µm-deep rectangular cross section.

The electrical conductivity of the fluid is assumed to be uniform throughout the T-junction domain. In addition, we assume that the analyte concentration is sufficiently low so as to not alter the conductivity of the solution during the injection process. The electroosmotic flow is assumed to be zero to simplify the simulation. For the hydrodynamic flow, it is assumed that the viscosity of the solution is not altered by presence of the analyte, which is generally valid for small molecules at low concentration. We also assume no pressure-driven flow in the separation arm, L3. Under these assumptions, the equations governing the electric potential and the hydrodynamics are not coupled to each other or the concentration of the analyte. For the hydrodynamic flow, we use zero velocity (no-slip) boundary conditions on the channel walls, and apply pressure boundary conditions at F1, F2, and F3. A pressure-driven flow between F1 and F2 may be established by setting P1 ) P, P2 ) 0, and P3 ) P/2. The corresponding flow rates Qi at each of the faces are Q1 ) -Q2 and Q3 ) 0, or all of the fluid entering the domain through F1 exits through F2. Similarly, an electric field may be established between faces 2 and 3 by applying V2 ) V, V3 ) 0, and V1 ) V/2. Thus, species transport in the T-junction will be purely pressure-driven between F1 and the T-junction and purely electrophoretically driven between the T-junction and F3. The advection-diffusion problem in the aforementioned geometry was solved using FIDAP, a commercially available CFD package based on the finite element method. The solution methodology consisted of three steps: (1) solve for the steadystate hydrodynamic flow, (2) solve for the steady-state electric potential, and (3) solve for the time-dependent concentration of an injected analyte subject to the steady fields computed in steps 1 and 2. As expected, gradients in the pressure exist between faces F1 and F2, but no gradient exists between the T-junction and F3. Similarly, gradients in the electric potential exist between faces F2 and F3, but the region between F1 and the T-junction remains field-free. Analyte injection into the T-junction domain was performed by applying a time-dependent species concentration C1 at the inlet face F1. The form of the time-dependent boundary condition was a Gaussian pulse of 0.3-s duration, having a standard deviation of 0.05 s. Depending on the ratio of the average pressure-driven velocity vjp to the average electrophoretic velocity vjep, the sample can be either transferred completely to channel L2 or L3 or split between L2 and L3. For the case of a neutral species injected into the T-junction geometry, it is expected that the analyte transport

Figure 2. Injection of a neutral analyte into the T-junction geometry. Time sequence progresses from left to right and top to bottom. The elapsed time between successive images is 0.12 s.

will follow the stream lines of the pressure-driven flow between faces F1 and F2. The pressure boundary conditions at each of the three faces, F1, F2, and F3, were adjusted so as to achieve an average pressure-driven velocity of 1 mm/s, and the diffusivity of the analyte was assumed to be 3 × 10-6 cm2/s. Figure 2 shows a sequence of images that depict the concentration of the injected analyte in the T-junction as a function of time. The sample enters the domain through F1, travels through channel L1, and turns the 90° corner into channel section L2, eventually exiting the domain through face F2. A very small portion of the sample diffuses into the stagnant channel L3 while the bulk of the sample is transferred from L1 to L2. Band broadening due primarily to Taylor-Aris dispersion is evident from the rapid decrease in concentration of the injected plug, and the characteristic curved shape of the leading and trailing edge of the sample. For a charged analyte, we first examine the case in which vjep > vjp. In particular, we keep the average pressure-driven velocity vjp fixed at 1 mm/s and alter the analyte mobility such that average electrophoretic velocity is twice as large, or vjep) 2 mm/s. Figure 3 shows the injection sequence for this case, in which the analyte is completely transferred between channel segments L1 and L3. Conservation of the analyte flux at the T-junction requires the concentration of the analyte to be one-half as large in channel segment L3 as compared to L1, since C3vjep ) C1vjp. Substituting vjep ) 2vjp gives the dilution condition present at the T-junction, C3 ) C1/2. The dilution of the analyte is evident from the simulations in channel segment L3, in which the analyte is compressed to

occupy one-half of the channel and is spatially elongated. In this case, none of the analyte enters channel segment L2, because the electrophoretic velocity is always locally greater than the pressure-driven velocity. If the electrophoretic velocity of the analyte is much slower than the pressure-driven velocity, the analytes will split between channels L2 and L3. The injection sequence for this case is shown in Figure 4, in which the average pressure-driven velocity is twice the average electrophoretic velocity, that is, vjp ) 1 mm/s and vjep ) 0.5 mm/s. The injected analyte is equally split after the T-junction between channels L2 and L3. The injection sequence clearly demonstrates the difference in band broadening characteristics for the two flow modes. In channel segment L2, the band broadening is dominated by flow-induced Taylor-Aris dispersion, whereas in segment L3, the band broadening occurs via molecular diffusion of the analyte. EXPERIMENTAL SECTION Apparatus. All experiments were performed either with a homemade development station or a modified Caliper 220 highthroughput screening system that was equipped with a multiport cartridge (Caliper Technologies Corp., Mountain View, CA). Caliper 220 systems are designed to provide a complete, integrated solution for primary assay screening and include automated sampling robotics with a complete software package for control and analysis. Three laser excitation sources with wavelength of 355, 451, and 633 nm, respectively, were used in Caliper 220s. Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

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Figure 3. Injection of a positively charged analyte into the T-junction geometry in which the average electrophoretic velocity is twice as large as the average pressure-driven velocity, that is, vjep ) 2vjp. The time sequence progresses from left to right and top to bottom. The elapsed time between successive images is 0.12 s.

The fluorescence from fixed locations on a microfluidic chip is monitored by three CCDs set to an emission wavelength band of 440, 530, and 685 nm, respectively, by band-pass filters. On the other hand, the development station was equipped with an arc lamp excitation source and PMT detectors. It is more flexible and can be set to monitor fluorescence from almost any location on a chip. It also consists of a video imaging system that allows one to visually inspect and observe the flow pattern. However, the detection sensitivity in the development station is ∼2 orders of magnitude poorer than in the laser-based Caliper 220 systems. The chip mounts inside a cartridge, which provides the interface and alignment to the multiport pressure and voltage controller. The design of the multiport control module was published previously.11 Briefly, the multiport control module provides basic control capabilities needed for microfluidic chips. Using ambient air as the control medium, eight independent peristaltic pumps can provide 5 psi of either positive or negative (vacuum) pressure. The voltage controller provides eight separate high-voltage lines capable of reaching ( 3 KV. All firmware and data collection software was designed in-house and runs on a PC. The multiport module is typically controlled through a script that contains the order, duration, and magnitude of each function, such as the pressure or voltage settings. Microchip Fabrication. Microfluidic devices were designed to have channels of known resistance through precise control of channel length, width, and depth. Standard semiconductor pho5178 Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

tolithography processes were used to etch channels, as reported previously.19 A quartz plate was drilled with an array of eight 2-mm holes for reagents. Photomasks of the devices were used to print the channel layout onto photoresist spun over quartz glass surfaces. Controlled hydrofluoric etching determined the channel depth. Holes for capillary tubes used for sipping compounds onto the chip were drilled into a separate quartz plate. Closed microfluidic conduits were created by aligning the two plates and bonding them together by a high temperature thermal process, as described by several authors.20-22 Finally, a capillary tube was inserted perpendicular to the planar surface of the chip to create a sipper chip. Chip Description. The schematic diagram of the microchips used in our experiments is shown in Figure 5. This chip consists of a central main channel in which the sipper attaches at one end and the other end terminates in a waste well. Two side electrode channels, separated by 31.5 mm, intersected the main channel. The chip was originally designed for performing an electrophoretic separation in high-throughput operation where the flow is driven by a single vacuum source at the far end of the chip. The chips (19) Kerby, M.; Chien, R.-L. Electrophoresis 2001, 22, 3916-3923. (20) Mathies, R. A.; Simpson, P. C.; Woolley, A. T. In Micro Total Analysis System ’98; Harrison, D. J., van den Berg, A., Eds.; Kluwer Academic Publishers: dordrecht, The Netherlands, 1998; pp 1-6. (21) Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1996, 68, 720-723. (22) Schmalzing, D.; Koutny, L.; Adourian, A.; Belgrader, P.; Matsudaira, P.; Ehrlich, D. Proc. Natl. Acad. Sci. U.S.A. 1998, 94, 10273.

Figure 4. Injection of a positively charged analyte into the T-junction geometry in which the average pressure-driven velocity is twice as large as the average electrophoretic velocity, that is, vjp ) 2vjep. The time sequence progresses from left to right and top to bottom. The elapsed time between successive images is 0.12 s. Table 1. Geometric Configurations and the Hydrodynamic Resistances of the Channels

Figure 5. Schematic diagram of the chip design. The geometric configuration of the channels and the predicted hydrodynamic resistances are listed in Table 1. The detector location for the development station and the Caliper 220 system are marked as D1 and D2, respectively. The SIE separation is achieved in the T-junction between channels R1, R2, and R3.

were modified to have a single depth of 10 µm in all channels. The geometric configuration of the channels and the predicted hydrodynamic resistances are listed in Table 1. The viscosity of the solution was assumed to be 1 centipoise. The detector

channel

length (mm)

depth (µm)

width (µm)

resistance (g/cm4 s)

sipper R1 R2 R3 R4 R5

20 13 16.6 31.5 13.05 13.76

10 10 10 10 10

20 25 210 45 270 70

1.13 × 1011 1.00 × 1010 1.12 × 1011 6.06 × 109 2.81 × 1010 5.09 × 1010

locations for the development station and the Caliper 220 system are marked as D1 and D2, respectively. Using a multiport module capable of providing multiple pressure and voltage control, we can design a flow pattern such that the bulk solution and neutral substrates flow from the sipper down the first electrode sidearm accompanied by a small backflow in the separation channel from the second electrode well. As we have shown previously, precise flow control from each individual channel can be achieved,assuming that the hydrodynamic resistance of the network is known.11 When high voltage is applied across the separation channel, the electric field produces electrokinetic forces that are superimposed on the pressure-driven forces. Analytes with sufficient electrophoretic mobility escape this separation junction and flow downstream past the detector to the second electrode. This technique can be applied to systems either with or without electroosmotic flow. Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

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The Caliper 220 analysis systems hold the chip such that the fluorescence detection region is located close to the end of the separation channel but prior to the downstream electrode. The detection location for the development system is usually set at 2 mm away from the T-junction, as shown in Figure 5. This assay chip was chosen simply to demonstrate the concept of selective ion extraction of one charged compound from a mixture of compounds. A chip designed to optimize the separation process according to SIE principle with a much shorter distance to the detection point is in progress. Reagent Preparation. Deionized water (18.2 MΩ-cm at 25 °C) used to prepare reagents was purified using a Milli-Q system. A 1 M HEPES buffer at pH 7.5 was prepared using ULTROL grade HEPES in both the free acid (Calbiochem, San Diego, CA) and sodium salt form (Calbiochem). All solutions were filtered through 0.2-µm polypropylene syringe filters before addition to the chip. Aqueous solutions of a peptide substrate and product specific for protein kinase A were prepared in an assay buffer at pH 7.5. Protein kinase A (PKA), a cAMP-dependent protein kinase, (Promega, Madison, WI) was reacted with a custom-synthesized substrate 5-FAM-LRRASLG-CONH2 (Caliper Technologies Corp., Mountain View, CA) of molecular weight 1129.5. The 5-FAM is a fluorescein NHS ester moiety (Molecular Probes, Eugene, OR) attached to leucine on the amino terminus of the peptide. The custom peptide purity was g98%, as measured by HPLC. The PKA assay buffer consisted of 5 mM MgCl2 (Sigma, St. Louis, MO), 0.01% Triton-X (Sigma), 1 mM DTT (Calbiochem), 10 µM ATP (Sigma), and 2% DMSO (Burdick & Jackson, Muskegon, MI) in 100 mM HEPES buffer. Dynamic coating reagent 3 (Caliper Technologies Corp.) was added to the buffer to suppress electroosmotic flow. At pH 7.5, the PKA enzyme converts the neutraly charged (Z ) 0) substrate into a negatively charged (Z ) -2) product. Aliquots of enzyme and substrate stock solutions were stored at -80 °C until needed. All solutions were stored on ice prior to reaction. In a polypropylene centrifuge tube, 100 µL of assay buffer containing 100 µM substrate and 25 nM enzyme were allowed to react to completion at room temperature for 90 min. The assay buffer was filtered at 0.2 µM prior to addition of the enzyme and substrate and 80 µL of 10 mM EDTA (Sigma) was added to stop the reaction. The purity of the product was checked via capillary electrophoresis, and the concentration was verified via UV absorption using an extinction coefficient () of 82 000 M-1cm-1 at 508 nm. Aliquots of product and substrate were stored at -80 °C until needed for individual experiments. RESULTS AND DISCUSSION Window of Separation. There are many parameters, voltages, or pressures that could be varied to achieve selective ion extraction. As a simple proof-of-concept experiment, the pressure on the sidearm (P2) was varied while keeping all other parameters fixed. If P2 is low enough, both neutral and charged compounds injected from the input channel (L1) will flow into the side channel (L2), as shown in Figure 6a. There is also a backflow (Q3) in the output channel (L3) toward the T-junction and into the side channel. In this case, the optical detector located in L3 should detect a steady background signal only. As P2 gradually increases toward P3, the hydrodynamic velocity in L3 decreases while the electrophoretic velocity remains constant. The total velocity in the output channel will reverse direction, and the compounds are 5180 Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

Figure 6. Schematic diagrams of three different flow patterns as the pressure on the sidearm varies. A constant voltage (V3 - V2) is applied in all cases. The detector is located on the lower left arm (L3) of the T-junction. (A) Reverse pressure-driven velocity in the output arm (L3) is greater than the forward electrophoretic velocity of the species resulting in no species detected. (Q3 is negative) (B) Reverse pressure-driven velocity in the output arm is less than the forward electrophoretic velocity of the species, resulting in species detected. (Q3 is positive) (C) A forward pressure-driven velocity in the output arm and sidearm plus the forward electrophoretic velocity of the species results in species detected and diluted. (Q3 is positive).

extracted at some predetermined pressure setting, as shown in Figure 6b. The exact pressure setting for the transition between the flow patterns shown in Figure 6a and b depends on the electrophoretic mobility of the analytes; the compound with highest electrophoretic mobility is extracted first. For a range of pressure settings, the analytes from the input channel L1 will split after the T-junction between output channels L2 and L3, as described in the previous section. In a system in which the chip is prepared using a uniform buffer with a constant concentration of background electrolytes, the distribution of electric field strength will remain constant if the concentration of analytes is much smaller, as compared with background electrolytes. An analyte plug extracted from the junction, hence, retains the same concentration but with different velocities that are determined by the combination of hydrodynamic and electrokinetic forces. Neglecting the slightly broadened dispersion occurring in the junction, the optical detector should yield a constant signal regardless of pressure variation in the external sources for this second case of flow pattern shown in Figure 6b.

Figure 7. (a) Flux model of PKA product (dotted line) and substrate (solid line) concentration at the detector as a function of pressure on the side well. This plot is specific for both the chip used and the voltage applied. Modeled at 750 V/cm. (b) Experimental data for PKA product (dotted line) and substrate (solid line) concentration at the detector. E-field set to 750 V/cm.

Finally, as P2 further increases, the flow direction in the side channel (L2) reverses and results in a dilution of the compound from the input channel (L1), as shown in Figure 6c. Neutral species or slower ions will extract into L3 eventually. In this case, the concentration of the analytes, hence the optical signal, will drop as the result of dilution. A simple species flux and flow velocity calculation was used to predict the separation of PKA substrate (Z ) 0) and product (Z ) -2) peptides as shown in Figure 7a. For a mixed sample of product and substrate flowing down the input channel, a sidearm pressure of -0.58 psi, both the substrate and product reach the detector, partially diluted by inflows from the sidearm. To test the model, a range of the sidearm pressures was slowly scanned while continuously sipping a peptide. The experimental results are shown in Figure 7b. Beginning at -2 psi on the sidearm, the pressure was increased every 200 s while sipping the peptide in a buffer-filled chip. The fluorescence intensity at the detector was recorded prior to taking the next pressure step. These intensities were normalized and plotted as concentrations for both product and substrate. Since the mechanism of SIE is a summation of hydrodynamic and electrophoretic velocities, we expect that at the transition boundaries, the peptides have a net forward velocity close to 0 as they pass the detector. This experiment was performed in a Caliper 220 system in which the detector is located 17 mm away from the separation junction. This long distance to detection, and therefore, a time delay, resulted in a slight shift of the concentration curves in Figure 7b toward higher sidearm pressures. Overall, the agreement between the theoretical curves predicted and experimental data is very good. The experiment described above shows a window of separation between two compounds with different electrophoretic mobilities as the pressure on the side channel changes. However, when running an assay, the pressure settings are usually fixed to maintain a constant flow rate and consistent reaction time. Nevertheless, the model and experimental results demonstrate the ability to selectively detect only one species with a relatively constant optical signal within a wide range of pressure settings. This feature of constant signals regardless of the pressure fluctuation is one of the main advantages of SIE over other conventional separation methods. Plug Separation. To evaluate the SIE model with short fluid plugs containing peptides, four wells of a 384-well plate were filled to contain PKA buffer, 1 µM PKA substrate, 1 µM PKA product, and a mixture containing 1 µM each of substrate and product, respectively. The chip was filled with PKA buffer containing a dynamic coating and loaded into the multiport Caliper 100 system. Using micrometers attached to the optical system, we were able to reproducibly move the detector to any location on the chip. A video camera and monitor connected to the optical system allowed for visualization of the detector location on chip. The detector was first located 0.2 mm prior to the separation junction, using the inner corner of the T-junction as a reference point. On the basis of the model, the pressures and voltages applied to the chip established conditions such that no product or substrate was expected to pass the separation junction. Next, the robot moved the well plate to introduce sample plugs from the plate to the sipper. Including the latency of robot motion, sample plugs of 1.7-s duration were spaced by 60 s of buffer sipping. A series of peaks were collected at the detector, which provided the prejunction peak characteristics. Next, the detector was repositioned 0.5 mm from the inner corner of the T-junction where the SIE junction ends. The sampling of the well plate was repeated to collect the postjunction data. Figure 8 summarizes the results that demonstrate three regions of operation using SIE of PKA compounds recorded before and after the tee separation junction. The fluorescence peaks are labeled as B, S, P, and S + P corresponding Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

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Figure 8. The three regimes of operation. Fluorescence intensity peaks of PKA compounds recorded before and after the tee separation junction: B, buffer; S, substrate; P, product; S + P, product and substrate. Chip end pressure is constant at 0.5 psi. Voltage is constant at 2800 V (a) Side pressure (P2) is set at -1.5 psi. The pressuredriven velocity in the detector channel is -1.2 mm/sec, which is opposite the forward electrophoretic velocity of product at 0.78 mm/ sec. As expected, no peaks reach the detector. (b) P2 ) -1.0 psi. At the reduced pressure, reverse flow slows to -0.59 mm/sec against forward electrophoretic flow of 0.78 mm/sec. Only product reaches the detector. (c) P2 ) -0.5 psi. In the detector channel, the pressuredriven flow is slightly positive at 0.12 mm/sec, and electrophoretic flow remains at 0.78 mm/sec. Both product and substrate are detected. The slowly traveling substrate peaks have broadened as a result of dispersion. 5182 Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

to buffer, substrate, product, and product and substrate, respectively. The small peaks between samples in the figure are optical noise due to the robotic movement in our prototype machine that is not present in Caliper 220. The three sidearm pressures used, (-1.5, -1, and -0.5 psi) fall in selection windows for nothing detected, product only detected, and both detected, respectively. Since we can adjust the position of the optics much closer to the separation junction, the travel time of signal peaks is not subject to the same time delays as we reported in the previous section. Ideally, within the product selection window, the chip should be operated at a sidearm pressure close to the substrate breakout point where the forward velocity of the product is greatest. Again, the simple flux model is a useful tool for predicting SIE voltage and pressure requirements. Dispersion. One concern about SIE is whether excessive dispersion results from the extraction junction design. Using the same short-plug dataset collected above, dispersion caused by crossing the junction was evaluated. This set also included data collected when the detector was repositioned a third time at 1 mm past the junction. The full width at half-maximum (fwhm) values were determined using the peak-fitting module in Origin 6.0 (OriginLab, Northampton, MA) software. Prior to passing the separation junction, the fwhm of the peaks was 2.57 s. At 0.5 mm after the junction, the fwhm average of the peaks (n ) 3) was 3.02 s. At 1 mm following the junction, the fwhm (n ) 3) was 3.15 s. The fwhm values before and 0.5 mm after the junction show an increase of only 17.3%. At 1 mm, the fwhm values increase to 22.3% above the prejunction values. This result is pleasing, considering the lack of symmetry in the T-junction of this chip. The channels are uniformly deep at 10 µm but nonsymmetric in width. The input, side and output arms are 25 µm, 200 µm, and 45 µm, respectively. Taylor dispersion appears to be the most likely contributor to the increase in fwhm values. As the vacuum on the side well is decreased, the reverse flow in the output arm decreases. The velocity of the traveling peak is the vector sum of the electrophoretic velocity minus the hydrodynamic velocity in the reverse direction. As the vacuum on the sidearm decreases and the backflow in the output arm decreases, the net forward velocity of the product plug increases. The nondispersive plug flow profile of electrophoretic flow predominates over the dispersive parabolic hydrodynamic flow. As hydrodynamic flow decreases, the dispersive effects of the reverse flow decreas, and we therefore see smaller changes in the fwhm on either side of the T-junction. The dispersion could be minimized further by redesigning a chip specifically for SIE. One of the major applications of SIE is high-throughput screening (HTS). To demonstrate the speed advantage provided for HTS, we performed a rapid sipping experiment designed to minimize the cycle time between sample sips yet maintain a high resolution. To prevent mixing of successive sample plugs by diffusion and dispersion, each sample is separated using a spacer of buffer, which we seek to minimize. The experiment was performed on a multiport Caliper 100 system, on which the detector region was positioned 0.5 mm away from the separation junction. A mixture of 1 µM PKA substrate and product were sipped as 1.7-s-long plugs onto the chip. Starting with a buffer sip time of 7.5 s, the buffer sip time was reduced to 3.5 s in 1-s

Figure 9. Rapid separation of product from five sample plugs of 1 mM product and substrate. Each plug of 1.7-s duration was spaced by 3.5 s of buffer. The buffer and sample times include 1.5 s of travel time required by the robot to change samples. To establish SIE operation for product, the pressures were -0.5 psi on the second electrode and the waste wells and -1 psi on the first electrode well, while the sipper remained open to the atmosphere.

increments. Individual peaks were easily discernible and maintained at a resolution >1 when utilizing a sip interval as short as 5.2 s, as shown in Figure 9. Separation Efficiency. To demonstrate the power and efficiency of separation of SIE, we mixed a series of PKA substrate and product mixtures with different ratios from 10:1 to 1000:1, which simulated low conversions of product in a high background substrate signal. Figure 10a shows the electropherogram produced by the conventional flow configuration where both substrate and product flow down to the detector. The large background substrate signal masks the product signal peaks completely until the product comprises 4% of the mixture. Even so, the analytical relevance of this peak is questionable. Even for products with reasonable electrophoretic mobility shift, a long migration time in the separation channel is needed to achieve adequate separation power. Figure 10b presents data from an SIE configuration using the same peptide mixtures in which only the product is allowed to migrate to the detector. The small fraction of selected product is easily detected out of the large background signal. The detection limits obtained from these figures are about 2 nM. This sensitivity could be improved even further if the detection location was closer to the separation junction to prevent unnecessary dispersion in the separation channel. CONCLUSIONS It is relatively easy to get baseline separation between a -2and 0-charged peptides in capillary electrophoresis. However, it is a challenge to convert it into high-throughput format on microchips. We have demonstrated a new separation technique, which relies on the combination control of multiple pressure and voltages sources that is ideally suited for microfluidic devices. A simple velocity balance model proved useful for determining the voltage and pressure settings used to predict separation. Using multiport control, the flow in any section of the microfluidic network can be configured to selectively extract the desired analytes on the basis of electrophoretic mobility. Although a simple T-junction was used in this manuscript, the same technique can

Figure 10. Test for separation sensitivity to low PKA product concentrations, shown as percentage, mixed in 1 µM substrate. A 1% peak represents 10 nM product. (a) Test for sensitivity in a conventional off-chip assay in which both substrate and product move past the detector. Product peaks arrive before the larger substrate peaks. The large background substrate signal masks the product signal peaks completely until the product comprises 4% of the mixture. (b) Sensitivity test under SIE operation for product. Peaks shown represent product that was separated from substrate. No substrate peak was observed.

easily be extended to a cross-junction or other complicated channel network. We have successfully extracted several enzymatic products of varying mobilities (positive, negative, and neutral) from their unmodified substrate pairs. SIE produces a highly efficient separation while adding minimal dispersion. This technique is well-suited for a high-throughput screening system. ACKNOWLEDGMENT The authors thank Andrea Chow and Theo Nikiforov for many inspiring discussions. We thank Bahram Fathollahi and Hugh Daniels for providing us chemicals, mobility data, and support for the experiment and simulation. Received for review May 28, 2002. Accepted August 14, 2002. AC0258103 Analytical Chemistry, Vol. 74, No. 20, October 15, 2002

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