Anal. Chem. 2000, 72, 3745-3751
Generation of Natural pH Gradients in Microfluidic Channels for Use in Isoelectric Focusing Katerˇina Macounova´, Catherine R. Cabrera, Mark R. Holl, and Paul Yager
Department of Bioengineering, Box 352141, University of Washington, Seattle, Washington 98195
As a part of an ongoing investigation of the use of isoelectric focusing (IEF) in microfluidic devices, pH gradients were electrochemically formed and optically quantified in microfluidic channels using acid-base indicators. The microchannels consisted of two parallel 40-mm-long electrodes with an interelectrode gap of 2.54 mm; top and bottom transparent windows were separated by 0.2 mm. Gradients in pH were formed as a result of the electrochemical decomposition of water at an applied potential not higher than 2.5 V to avoid generation of gas bubbles. Solutions contained low concentrations of a single buffer. The stability of the pH gradients and their sensitivity to changes in initial conditions were investigated under static (nonflow) conditions. Isoelectric focusing of sample biological analytes, bovine hemoglobin and bovine serum albumin, was performed to illustrate the potential of “microfluidic transverse IEF” for use in continuous concentration and separation systems.
A preconditioning system for the detection of chemical and biological warfare (CBW) agents being developed in our laboratory consists of three sequential processes. In each process, a force is applied to analytes transverse to the flow in microchannels. The first step, continuous separation of particles on the basis of variation in sedimentation rate, has been performed and will be described elsewhere.7 Analyte particles will be separated from interferent particles and concentrated in two subsequent continuous processes using electrokinetic-based techniquesszone electrophoresis and isoelectric focusing (IEF). Zone electrophoresis is today commonly performed as a batch chromatographic technique in capillaries (capillary electrophoresis) and in microfabricated devices.8,9 In this method, hydrated charged particles are separated in an electric field according to their electrophoretic mobility (µe)
Microfabrication and miniaturization of analytical instrumentation1,2 have been extensively investigated during the last 10 years. Such microfabricated systems offer many potential advantages over conventional analytical systems, such as low sample consumption, lower device cost, reduced waste creation, and, possibly, reduced analysis time.2-6 These devices will enable novel investigative methods in many areas including biochemistry, clinical chemistry, and agrochemical research. To expand the utility of microfluidic technology, these devices must be designed to function with “real world” samples. When microfluidic devices are used with samples such as blood or the output from an air sampler, microchannel blockage by large particles or aggregates is a significant concern. A preconditioning system of appropriate design can prevent device blockage and ensure detection of analytes of interest without interference from other irrelevant compounds or particles.
which is a function of ion charge (q), ion radius (r), and fluid viscosity (η). Higher resolution and lower band diffusion can be achieved by isoelectric focusing (IEF) performed in pre-established pH gradients;10 the species are separated according to their isoelectric points (pI). A molecule or larger particle migrates electrophoretically through the pH gradient until it reaches the position in the pH gradient that corresponds to its pI, at which point the net molecular charge on the particle is zero.11 While initially performed in macroscopic devices,12 IEF today is usually performed in devices with at least one dimension less than 1 mm, such as fused silica capillaries.10,13 The distance between the electrodes is generally large, as is the applied field. Often the capillaries are filled with a polymeric gel to minimize convective disturbances. The IEF technique has recently been performed in microfabricated devices.5,14 Microchannels in a glass wafer were fabricated using photolithography and chemical etching.14 A voltage of 3 kV was
* Corresponding author. E-mail:
[email protected]. (1) Fintschenko, Y.; vandenBerg, A. J. Chromatogr., A 1998, 819, 3-12. (2) Qin, D.; Xia, Y. N.; Rogers, J. A.; Jackman, R. J.; Zhao, X. M.; Whitesides, G. M. Microsystem Technology In Chemistry And Life Science 1998, 194, 1-20. (3) Brody, J.; Yager, P.; Goldstein, R.; Austin, R. Biophys. J. 1996, 71, 343041. (4) Brody, J.; Yager, P. Sens. Actuators, A 1997, A58, 13-18. (5) Hofmann, O.; Che, D. P.; Cruickshank, K. A.; Muller, U. R. Anal. Chem. 1999, 71, 678-686. (6) Rossier, J. S.; Schwarz, A.; Reymond, F.; Ferrigno, R.; Bianchi, F.; Girault, H. H. Electrophoresis 1999, 20, 727-731. 10.1021/ac000237d CCC: $19.00 Published on Web 06/30/2000
© 2000 American Chemical Society
µe )
q 6πηr
(1)
(7) Holl, M. R.; Macounova, K.; R., C. C.; Yager, P. Electrophoresis, submitted for publication. (8) Effenhauser, C. S.; Bruin, G. J. M.; Paulus, A.; Ehrat, M. Anal. Chem. 1997, 69, 3451-3457. (9) Li, P. C. H.; Harrison, D. J. Anal. Chem. 1997, 69, 1564-1568. (10) Kane, M.; Nishimura, A.; Nishi, K. Anal. Chim. Acta 1999, 383, 157-168. (11) Andrews, A., T. Electrophoresis: theory, techniques and biochemical and clinical applications; Clarendon Press: Oxford, 1986. (12) Svensson, H. Acta Chem. Scand. 1961, 15, 325-41. (13) Righetti, P. G.; Bossi, A. Anal. Chim. Acta 1998, 372, 1-19. (14) Mao, Q. L.; Pawliszyn, J. Analyst (Cambridge, U.K.) 1999, 124, 637-41.
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applied across electrodes placed at the ends of the microchannel. As with all such high-voltage applications, the generation of O2 and H2 by electrolysis required that the electrodes be vented. Application of the electric field perpendicular to a flowing fluid stream would enable continuous separation of particles on the basis of their size and charge. In 1983, Giddings and colleagues first proposed the use of an electric field in combination with field flow fractionation (EFFF).15 Simultaneous use of an electric field and a pH gradient formed in a device could be used for separation of biological particles on the basis of pI. Chmelik16 and Thormann17 first published experimental data demonstrating the use of such a system to fractionate a binary protein mixture. Devices with electrode gaps ranging from 0.2 to 12 mm were operated in batch mode only. The pH gradients were formed using carrier ampholytes; the electrodes required large buffer reservoirs with cooling and recirculation. The use of transverse IEF, as defined by Chmelik16 and Thormann,17 has many advantages over that of traditional IEF. However, several problems remain; removal of heat generated by Joule heating, convective disturbance of the sample stream that decreases resolution, and the requirement for high voltages to generate sufficient field strength are among the main problems. Microchannels formed by two long parallel electrodes producing an electric field perpendicular to the flow direction of the fluid offer several improvements: (1) generation of high field strength at lower absolute voltages at the electrodes due to a small interelectrode gap, which could reduce gas generation; (2) a high surface-to-volume ratio that facilitates heat transport, thus reducing or even eliminating the need for cooling; and (3) minimization of convective disturbances within the separation chamber due to both laminar flow and minimal heating. A key feature of the proposed microchannels is the absence of electrolyte reservoirs and integration of the electrodes with the walls of the separation channel. By having the electrodes in intimate contact with the separation channel, the products of electrolytic decomposition of water can be used as a source of H+ and OH-. The pH gradients are rapidly formed as a result of the electrolysis of water.18 Oxidation of water takes place at the anodic surface, forming H+ and O2 gas (eq 2). Reduction of water at the cathode leads to formation of H2 gas and OH- (eq 3).18
Anode: H2O S 2H+ + 1/2O2 + 2e-
(2)
Cathode: 2H2O + 2e- S 2OH- + H2
(3)
The maximum possible applied voltage is limited by the need to avoid generation of gas bubbles, which would interfere with uniform flow in the microchannel. In situ detection of pH gradient formation is an experimental challenge due to the extremely small fluid volumes being processed and the spatial and temporal variation of pH within the (15) Giddings, J.; Lin, H.; Caldwell, K.; Myers, M. Sep. Sci. Technol. 1983, 18, 293-306. (16) Chmelik, J.; Deml, M.; Janca, J. Anal. Chem. 1989, 61, 912-14. (17) Thormann, W.; Firestone, M. A.; Dietz, M. L.; Cecconie, T.; Mosher, R. A. J. Chromatogr., A 1989, 461, 95-101. (18) Corstjens, H.; Billiet, H. A. H.; Frank, J.; Luyben, K. C. A. M. Electrophoresis 1996, 17, 137-43.
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channel. Amphoteric dyes have been used previously as low molecular weight markers for spectroscopic monitoring of pH changes in capillaries18 and channels.19 Acid-base indicators were chosen to monitor pH changes in our microfluidic channels. However, indicator dyes are not useful for characterizing shallow pH gradients, in part because of their broad titration range. Use of labeled proteins with known pI values that can be focused into a narrow band is preferred. Therefore, bovine hemoglobin (pI ) 7.1) and a fluorescent conjugate of bovine serum albumin with Bodipy FL (pI ) 4.65) were employed as sample analytes. Additionally, the use of proteins to monitor pH gradient formation allows us to demonstrate the potential utility of this technique for sample preconditioning. In the present work, microfluidic channels formed by two parallel gold electrodes are described.7 The development of pH gradients formed in these microchannels under static conditions was quantified using the acid-base indicators. It is shown that such microchannels can be used for IEF of proteins. EXPERIMENTAL SECTION Chemicals and Instrumentation. Acid-base indicators chemically derived from sulfopthalein were used to monitor pH gradient formation. Bromocresol purple (pKa ) 6.3) and phenol red (pKa ) 7.9) were used as purchased (Aldrich Chem. Co., Milwaukee, WI). The concentration of indicators in aqueous solutions was usually 0.2 mM. As the supporting electrolyte, Na2SO4 (Aldrich) was used. Histidine (Research Chemicals Ltd., Avocado, Lanc.) and 2-(4-morpholino)ethanesulfonic acid (MES; Fisher Biotech, Fair Lawn, N. J.) were employed as amphoteric buffers. Initial pH values were measured with a pH meter (Orion, model 290A). Bovine hemoglobin (Sigma Chem. Co., St. Louis, MO, pI ) 7.1) and fluorescent bovine serum albumin (BSA)/Bodipy FL conjugate (Molecular Probes, Inc., Eugene, OR) were used as test analytes for isoelectric focusing. The isoelectric points (pI) of proteins were determined experimentally by polyacrylamide gel isoelectric focusing. The gel IEF was carried out in carrier ampholyte Bio-LYTE with a pH range from 3 to 10 (BioRad Laboratories, Inc., Hercules, CA), using the standard procedure specified by BioRad Laboratories, Inc. A mini IEF cell (model 111) purchased from BioRad and equipped by two graphite electrodes, was employed for this purpose. Focusing was carried out under constant voltage conditions (Power Pac 1000, BioRad). Characterization of Protein Analytes. Isoelectric points of proteins were determined by comparison with the positions of standards (IEF Standards with a pI range from 4.45 to 9.60 (BioRad Laboratories, Inc., Hercules, CA)). Bovine hemoglobin showed multiple bands at a position close to that of standard human hemoglobin A (pI ) 7.1). The position of the fluorescent BSA conjugate band corresponded with the position of a second band of phycocyanin (pI ) 4.65). The stability of the fluorescence of the BSA conjugate was measured over a broad range of pH values with an LS-50B Perkin-Elmer luminescence spectrometer (Norwalk, CT). Electrochemical Flow Cell. An electrophoretic flow cell was fabricated in polyester (Mylar) using laser ablation micromachining methods described elsewhere.7 Figure 1A is an exploded view (19) Slais, K.; Friedl, Z. J. Chromatogr., A 1995, 695, 113-22.
Figure 2. pH changes of a 0.2 mM solution of phenol red in 1 mM histidine and 5 mM Na2SO4; applied potential of 2 V, current density of 0.25 A/m2, an initial pH of 7.6. (A) Optical images; ROI after application of 2 V to the wires to the electrodes. (B) Specific pH values of phenol red and bromocresol purple as determined from images; 9 specific pH locations for alkaline front of phenol red, 2 specific pH locations for acid front of phenol red, 4 specific pH locations for acid front of bromocresol purple (actual images not shown).
Figure 1. Polymeric laminate electrochemical flow cell. (A) Illustration in exploded view: (a) upper observation window with fluid vials for inlet and outlet; (b) upper cap; (c) flow cell end cap; (d) electrode substrate; (e) deposited gold layer; the Mylar onto which gold had been deposited is wrapped around a Mylar core to form the electrode; (f) lower cap; (g) observation window. Layers b and f are precoated on both sides with acrylate adhesive. The small holes seen on layers (c) and (d) were used for pin registration during the assembly. (B) Geometry and critical parameters, ROI (region of imaging).
of the flow cell. Laser micromachining was also used to create a lift-off mask for patterning of the gold electrodes. Gold (99.99% pure, Material Research Corporation) was deposited to a thickness of 0.1 µm on Mylar by rf sputtering without an intermediate adhesion layer. Activation of the Mylar surface with an O2 plasma prior to gold deposition produced a robust metal film. Individual flow cell components were assembled using an acrylate-based pressure-sensitive adhesive commonly used in the manufacture of disk drives (3M-1151). Mylar layers alternate with adhesives Mylar-adhesive layers. The register pins were used for easier assembly. The electrochemical flow cell channel geometry, critical electrode parameters, and the optical region of imaging (ROI) are illustrated in Figure 1B. The ROI is located in the middle of the channel to minimize electrode edge effects. An H-filter3,4 configuration flow cell was used with a main flow cell cross section of 0.41 mm between the two Mylar observation walls (z coordinate) and 2.54 mm between the two electrodes (y coordinate). The channel was 40 mm long (x coordinate). The two electrodes were 0.2 mm thick (along z coordinate) and centered between the top and bottom observation windows. In the x coordinate, the electrodes were 38.5 mm in length and centered between the inlet and outlet ports in the x coordinate. In all experiments, the cathode was located at the y-coordinate origin. Electrical connections to
the anode and cathode were achieved using silver epoxy or direct mechanical contact maintained by clamping. Visualization of the microfluidic channels was performed using an inverted optical microscope (IM 35, Zeiss, Germany). A lowpower objective (2.5/0.008) was used for all experiments. Images of the channel were taken using a 3-chip cooled CCD camera (ChromoCam 300, Oncor, Gaithersburg, MD) in combination with a video data acquisition card (CG-7 RGB frame grabber, Scion, Frederick, MD) and accompanying PC software (Scion Image). A standard fluorescein filter set (ex. 450-490 nm, dichroic at 510 nm, em. 520 nm long pass) was used for fluorescence measurements. Solution was injected into the channel through one of the inlets. If two inlets were employed, the channel was loaded with two syringe pumps (Kloehn Ltd., Las Vegas, NV) under computer control. In all experiments, flow along the x-axis was zero. For microfluidic IEF, a constant potential was applied between the gold electrodes by means of a dc power supply (model 612C, Hewlett-Packard, Palo Alto, CA) with an accuracy of 0.5%. Current values were measured by the same instrument. Measurement of current during the experiments and comparison with predicted current values has shown that there must be a large potential decrease somewhere in the IEF device, most likely immediately adjacent to the two electrodes. Therefore, the potential across the main body of the channel, calculated on the basis of experimentally measured current and conductivity values, was lower than 10% of the total voltage applied. Image Analysis. Data were obtained from optical images of the ROI in the channel. The ROI represented ∼4 mm of the channel along the x axis in all experiments. Three colors (dark red, moderately red, and yellow) were distinguishable in the channel filled by phenol red solution after a potential was applied (Figure 2A). Specific terminology was defined for easier description and understanding of experimental results. The boundary between two colors of the indicator dye is here referred to as a color front. In Figure 2A, the moderately red/dark red boundary represented the “alkaline front” (closer to the cathode); the yellow/moderately red boundary represented the “acid front” (closer to the anode). The position of the color front was Analytical Chemistry, Vol. 72, No. 16, August 15, 2000
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determined from a plot of green pixel intensity vs relative y position in the channel and was used to define the location of a specific pH. The color profile was measured along the field direction, and the “front” was considered to be a midpoint between a pH extreme (either acid or base) and the starting pH. For phenol red at an initial pH of 7.6 (36% deprotonated), these midpoints correspond to 18 and 68% deprotonated. For bromocresol purple at the same initial pH (100% deprotonated), there was only one visible color front corresponding to 50% deprotonation. From these deprotonation values, the specific pH was calculated using the HendersonHasselbach equation.The camera used in these experiments was determined experimentally to exhibit a log-linear response with respect to optical density (OD) (data not shown). That is, the log of the pixel value varies linearly with OD. By modeling total OD as the sum of contributions from the two forms of the indicator dye (each with a different absorption coefficient ), it can be shown that OD is linear with respect to the degree of deprotonation of the pH indicator dye
OD ) 1dCT + (1d + 2d)CTR
(4)
where d is optical path length, CT is total concentration of dye, R is the degree of deprotonation, and 1 and 2 are extinction coefficients of the protonated and deprotonated forms of the indicator. Therefore, the log of the pixel value also is linear with a degree of protonation. Experimental Conditions. Unless otherwise stated, all chemicals were used in these concentrations: 5 mM Na2SO4, 0.2 mM phenol red, 0.2 mM bromocresol purple, 1 mM histidine, 1 mM MES. The initial pH of solutions was adjusted using either 0.1 M NaOH or 40% (v/v) H2SO4. A constant voltage of 2.00 ( 0.01 V was applied throughout the experiments if Na2SO4 was used as the supporting electrolyte. After an initial transition period, (typically ∼20 s) in which the current dropped from 300 to 4 µA, the current remained constant. The applied voltage was chosen to maximize the electric field while avoiding production of gas bubbles. The applied voltage could be higher for solutions without Na2SO4. A potential of 2.5 V (5 µA) was used for IEF of hemoglobin, and a potential of 2.3 V (5 µA) was applied when BSA conjugate was focused. RESULTS AND DISCUSSION Use of Acid-Base Indicators. To illustrate the distribution of pH in the microchannel, two separate experiments were performed with different indicators (phenol red and bromocresol purple) at the same initial pH of 7.65. At this pH, phenol red is ∼50% deprotonated, resulting in an initial intermediate pale red color (Figure 2). Applying a potential of 2 V resulted in production of OH- at the cathode; the color in the vicinity of the electrode deepened to dark red. H+ was produced at the anode, causing the pale red color to fade to yellow at that side of the channel. As a result of a combination of electrophoresis and diffusion, the “alkaline front” and “acid front” moved toward each other until they reached “steady-state” positions (Figure 2A). It is possible to see that the steady state position was actually still slightly changing, even after 300 s. Therefore, the steady-state position was defined as the position of the “color front” when this position did not vary greater than 2% over a 5-min interval. 3748
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Bromocresol purple should be 99% deprotonated at pH 7.64 and in its basic purple form. When a potential of 2.0 V was applied, a change from purple to yellow was observed at the anode because of H+ production. The acid front moved toward the cathode over time until it reached a steady-state position. The specific pH locations for phenol red and bromocresol purple are shown in Figure 2B for each time increment. The numeric value of specific pH, calculated from the HendersonHasselbach equation, is posted at the right side of Figure 2B. These pH values describe the pH profile in the channel after the steady state was reached. Electrochemical Properties of Indicators. Acid-base indicators are electrochemically active compounds.20,21 Their use in monitoring the formation of pH gradients could be compromised if electrochemical reactions other than the electrolysis of water were to occur in the channel. Therefore, the electrochemical reduction of the indicator dyes was studied as a possible competitive reaction at the cathode (no oxidation of the indicator dyes was described in the literature). It is known that the structure of products after either electrochemical reduction or chemical reduction with sodium borohydride (NaBH4) are identical.22 Therefore, the chemical reduction of both phenol red and bromocresol purple with NaBH4 was examined as a model of electrochemical reduction. Histidine buffer NaBH4 (2 mg) was added to aqueous solutions of dyes (0.2 mM, pH 3.8) and incubated for 20 min. After 20 min, the characteristic color of the solutions disappeared, indicating that the chemical products were colorless. Since no comparable decolorization was observed at the cathode during any of our experiments, electrochemical reduction of the dye in the microfluidic channel under the conditions studied can be discounted. Effect of Initial Conditions on pH Gradient formation. Histidine Buffer. Sets of phenol red solutions in histidine buffer with different pH values with and without supporting electrolyte were employed. The formation of pH gradients was studied in unbuffered electrolyte solutions first. The pH gradients were established within two minutes at potentials of about 2 V, but their stability, especially for lower initial pH values, was poor. Better stability was observed for pH gradients developed in the presence of a buffer. Stable pH gradients developed in buffered electrolyte solutions in less than 5 min. Histidine with pKa values of 6.00 and 9.1723 was used as a buffer for the first experiments. It was found that the steady-state positions of the color fronts and, therefore, the pH distribution in the channel, were sensitive to the initial pH. The steady-state positions of the color fronts were established closer to the anode with increasing initial pH (Figure 3). Figure 3 demonstrates the difference in steady-state positions reached after 5 min for unbuffered and histidine-buffered electrolyte solutions. While the unbuffered solution (0.5 mM Na2SO4) only shows a gradual shift of steady state positions toward the anode with increasing initial pH, a plateau is seen for buffered solution (0.1 mM histidine in 0.5 mM Na2SO4). The plateau is seen for the initial pH range from 6.8 to 8.0 where buffer capacity (20) Kudirka, P. J. Anal. Chem. 1972, 44, 1786-94. (21) Prince, C. R.; Linkletter, J. G. S.; Dutton, P. L. Biochim. Biophys. Acta 1981, 625, 132-48. (22) Senne, J. K.; Marple, L. W. Anal. Chem. 1970, 42, 1147-50. (23) Bier, M.; Ostrem, J.; Marquez, R. B. Electrophoresis 1993, 14, 1011-18.
Figure 3. The effect of initial pH on pH gradient formation: (A) electrolyte solution (5 mM Na2SO4): change of position of steady state color-change fronts of phenol red with 2no buffer and with [ 1 mM histidine buffer; (B) 1 mM MES (without electrolyte): × dependence of steady state color-change fronts of phenol red on initial pH. Filled symbols illustrate different position of focused band of BSA conjugate for different initial pH after 2 5 min; 9 6 min; [ 10 min.
is weak (since that range is more than 1 pH unit from either pKa value for histidine). In this pH range, the majority of the buffer molecules are in their neutral form and therefore have minimal electrophoretic mobility (pI of histidine is 7.4). For IEF of proteins, solutions with no Na2SO4 were used. It was found by other authors that the presence of electrolytes significantly compresses pH gradients,24 which could affect the ability of microfluidic IEF as an efficient method for protein separation. However, no significant difference in the position of the color fronts of phenol red was observed between solutions with and without 0.5 mM Na2SO4 for initial pH values from 6.1 to 7.5. The absence of the additional electrolyte provided by the Na2SO4 decreased the conductivity, which increased the time needed for the pH gradient formation from 5 to 8 or 10 min. To decrease the development times, the applied potential was increased from 2.0 to 2.3 V. MES Buffer. Preliminary results showed that IEF of proteins worked best when the initial pH value of buffer solution was close to the pI of the protein. To achieve initial pH values close to the pI value of BSA, MES buffer (pKa) 6.09) was also examined. The dependence of the color-front positions on initial pH was investigated for 1 mM MES without electrolyte. As expected, the positions of the color fronts were closer to the anode for a higher initial pH. No plateau was observed on the curve (Figure 3B). The time needed for stable pH gradient development ranged from ∼8 min for a potential of 2 V (I ) 4 µA) and 4 min for a potential of 2.3 V (I ) 7 µA). Long-Term Acidification. If experiments with indicators (buffered solutions) were continued longer than 20 min (at an applied potential of 2 V), a slow movement of the acid and alkaline fronts toward the cathode was observed. This apparent acidification was not expected. Even though retention times of analytes in the IEF device of longer than 20 min are not expected in the microchannel under flow conditions, the cause of this slow acidification was still investigated. (24) Mao, Q. L.; Pawliszyn, J. J. Chromatogr., B 1999, 729, 355-9.
The apparent acidification could conceivably be due not to acidification, but to bleaching of the indicator dye by a high partial pressure of O2 generated at the anode. A high partial pressure of oxygen could also cause oxidation of target analytes in IEF of biological particles. To investigate the possibility of oxygen bleaching of indicators, the pH gradient formation protocol was modified; a solution of 0.2 mM phenol red in 0.1 M phosphate buffer at pH 10 was injected into the microchannel. At pH 10 the dye is strongly red, facilitating observation of bleaching. The high concentration of buffer should consume the electrochemically generated H+ and OH- ions, so no pH-mediated color change should be observed at the anode. However, if oxygen were to react with the indicator, a loss of red would accompany this reaction. No color change was observed at either electrode within 20 min, so chemical bleaching of indicator is not responsible for the slow movement of the color front of indicator migration toward the cathode. Drifting of the pH gradients is well-known in conventional IEF; several possible explanations have been suggested in the literature. Electroosmosis-induced mixing has been suggested25 as has isotachophoresis. Isotachophoresis is the electromigration of focused molecular bands in order of their mobility (and under certain circumstances can cause anodic drift).26 These explanations, as well as others, assume that stream fractions can leave the separation chamber and enter the electrolyte reservoirs via semipermeable membranes. Such an effect is not possible in our devices, since no electrolyte reservoirs are present. No adequate explanation has been found in the literature for the cathodic drift observed in our experiments. The possibility that this drift was caused by interactions between the solution and the “inert” materials of the device itself is still under investigation. Isoelectric Focusing. Bovine Hemoglobin. Having established that pH gradients were formed, the next step toward developing a practical continuous IEF microfluidic system was to focus a protein. Bovine hemoglobin (Hb) was selected as a test case because of its strong absorbance at 550 nm (facilitating observation without need for additional dyes). The pI of Hb is 7.1, as verified by polyacrylamide gel isoelectric focusing run at the same concentration as was used for IEF in the microchannel. IEF of Hb in the microchannel was performed in 1 mM histidine buffer with an initial pH of 7.1 (as was mentioned above, the best results of IEF of proteins were obtained for the initial pH of the buffer solution close to the pI value of the protein). A voltage of 2.5 V (resulting in a current of 5 µA) was applied for 6 min. Two zones of higher optical density close to the electrodes were formed within 15 s. They moved toward each other meeting to form one dark zone after 5 min. Even with a relatively high concentration of Hb (1.55 × 10-4 M), the darker zones and their migration were barely visible in color images. Normalized plots of green pixel values vs position allowed monitoring of Hb migration (Figure 4). The position of the final focused zone of Hb in the microchannel corresponded to a pH of 7.1 ( 0.3 on the basis of previous results with acid-base indicators under similar conditions (Figure 2). The pI value determined by IEF in the microchannel was, within experimental error, identical to that in the literature and that determined by our gel IEF experiment. (25) Hagedorn, R.; Fuhr, G. Electrophoresis 1990, 11, 281-9. (26) Mosher, R. A.; Thormann, W. Electrophoresis 1990, 11, 717-23.
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Figure 5. Actual images of the ROI for IEF of BSA conjugate in 1 mM MES buffer (no electrolyte); variation of the established band position with the different initial pH (E ) 2.3 V, I ) 5-8 µA).
Figure 4. Isoelectric focusing of bovine hemoglobin in 1 mM histidine buffer (no electrolyte, initial pH of 7.1, E ) 2.5 V, I ) 5 µA). The lines represent normalized color intensity profiles in the channel for different time steps.
A third absorbing zone formed near the cathode; it did not migrate but increased in intensity over time. This peak may be caused by electrochemical reduction of Hb27 or its adsorption at the gold surface.28 The presence of the secondary peak suggests that direct contact of proteins with the electrode surfaces is not desirable for practical IEF. BSA Conjugate. Because of its intense fluorescence and resistance to photobleaching, a commercially available conjugate of bovine serum albumin (BSA) with the fluorescent dye Bodipy FL allowed monitoring of IEF at lower protein concentration than was possible for Hb. A possible disadvantage of using such covalently modified proteins is the creation of a heterogeneous population because of variation in degree of conjugation with multiple labels. The pI of each type of conjugate could be different from the pI of the native protein because of the modification of the surface amines to other charged or neutral species. It has been shown that the conjugation reaction of BSA with other dyes, such as fluorescein isothiocyanate or rhodamine -B- isothiocyanate, does not necessarily significantly affect the pI of BSA (4.6).29 This conclusion was confirmed by our measurements of pI of the BSA conjugate via polyacrylamide gel isoelectric focusing; the pI was found to be 4.6 ( 0.1. The fluorescence of the BSA-Bodipy FL conjugate is reported to be insensitive to pH between 4 and 8.30 Since these pH values could be exceeded in the microchannels, particularly near the electrodes, the stability of the fluorescence signal of the BSA conjugate was studied, especially at higher pH values as found during the IEF. Solutions of 1.52 µM BSA were prepared for different pHs from 2.5 to 10.0. A fluorescence spectrum from 500 to 550 nm for an excitation wavelength of 495.0 ( 2.5 nm was (27) Zhang, S.; Sun, W. L.; Zhang, W.; Jin, L. T.; Yamamoto, K.; Tao, S. G.; Jin, J. Y. Anal. Lett. 1998, 31, 2159-2171. (28) Hook, F.; Rodahl, M.; Kasemo, B.; Brzezinski, P. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 12271-12276. (29) McDonagh, P. F.; Williams, S. K. Microvasc. Res. 1984, 27, 14-27. (30) Haugland, R. P. Handbook of Fluorescent Probes and Research Chemicals; Molecular Probes, Inc.: Eugene, OR, 1996.
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recorded for each of the solutions immediately after its preparation and then after each 30 min for 4.5 h (data are not shown). The fluorescence did not significantly change during these measurements for any pH values, confirming that the changes of fluorescence observed in the microchannels were not due to quenching by extremely low or high pH values. The IEF of the 2.9 µM BSA conjugate was performed in 1 mM MES buffer with an initial pH of 4.46 (if no OH- was added). No supporting electrolyte was used for these experiments. The channel was filled through two inlets. While a solution of buffer was loaded from the anode side, solution of BSA conjugate in the same buffer was loaded at the same speed from the cathode side. Flow was stopped immediately prior to application of the electric field. After applying a potential of 2.3 V (resulting in a current of 5 µA), the BSA conjugate ultimately focused into one tight band for all examined initial pH values. The position of the focused band, as well as time needed for focusing, varied for different initial pH values. As predicted by our previous investigation of pH gradient development, the BSA was focused closer to the anode for higher initial pHs (Figure 5). The positions of focused bands were assumed to match the pI of the BSA conjugate (pI ) 4.6) and were correlated with positions of color change fronts of phenol red measured under similar conditions (Figure 3B). The results were in good agreement and showed that shape of the pH gradient was not influenced by the presence of the protein. Time needed for IEF of BSA conjugate also varied with the initial pH. The IEF of BSA conjugate was faster for lower pH values. While IEF of BSA into one tight stream took place within 3 min for an initial pH of 3.54, 10 min was not long enough to focus the BSA conjugate into one tight band for an initial pH of 6.22. One explanation of this difference in time may be that the BSA conjugate had to travel farther at the higher initial pH values. The pH corresponding to the BSA conjugate pI shifted away from the cathode at higher initial pH values, which means shifting away from the initial position of BSA conjugate. CONCLUSIONS A method for characterization of pH gradients formed by electrolysis inside microchannels was developed. It was shown that natural gradients that are stable for many minutes can be produced in the microfluidic channels and that the pH profile depends on initial conditions. The utility of the microchannels for isoelectric focusing of proteins was demonstrated: bovine hemoglobin and a fluorescently labeled BSA were focused into single tight bands in a few minutes. The position of the focused protein bands was affected by the initial solution pH. Some important features were found that should be implemented in future designs of microchannels for flowing transverse
IEF. To avoid any direct contact of proteins with the electrodes, and thus avoid unpredictable behavior of protein at the electrode surface (as was observed in the case of IEF of Hb), three inlets could be utilized. The two outer streams that contact the electrodes would be free from protein. The proteins to be focused should be injected into the central stream. If gold electrodes are used, a potential lower than 2.5 V must be applied to avoid formation of bubbles. Changing the electrode material from gold to platinum or palladium, which are “nongassing” electrode materials, could help to overcome these problems.
ACKNOWLEDGMENT We would like to thank Prof. Wolfgang Thormann for his comments and helpful suggestions. We also would like to thank Andrew Evan Kamholz for valuable discussions. We are grateful to Mai Q. Nguyen and Olga G. Kaufman for help in performing experiments. This work was supported by DARPA Contract N660001-97-C-8632. Received for review February 25, 2000. Accepted May 23, 2000. AC000237D
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