Anal. Chem. 2005, 77, 6664-6670
Integrated Microfluidic Device for Solid-Phase Extraction Coupled to Micellar Electrokinetic Chromatography Separation Jeremy D. Ramsey†,§ and Greg E. Collins*,‡
Geo-Centers, Inc., Suite 200, Ballston Station, 4301 North Fairfax Drive, Arlington, Virginia 22201, and Chemistry Division, Naval Research Laboratory, 4555 Overlook Avenue, S.W., Code 6112, Washington, D.C. 20375-5342
An integrated microdevice was utilized for the autonomous coupling of solid-phase extraction (SPE) to micellar electrokinetic chromatography (MEKC). Porous plugs of polymethacrylate polymer (∼200 µm in length) were fabricated by ultraviolet irradiation in microchannels. Microcolumns of hydrophobic beads packed against the polymethacrylate plugs were utilized for the quantitative extraction of rhodamine B, yielding preconcentration factors over 200 for a 90-s extraction. The calculated detection limit for this dye was 60 fM. A sample of coumarin dyes were concentrated by SPE, eluted in a nonaqueous solvent from a separate on-chip reservoir, and injected by a gated valve onto a separate column for MEKC analysis. Using the integrated device, a completely automated sequence of extraction, elution, injection, separation, and detection were performed in less than 5 min. Observed separation efficiencies were high, with plate heights below 2 µm. The analysis was at least 3 times faster than semiautomated, conventional, solid-phase extraction, while requiring no user intervention. The design, fabrication, and autonomous operation of the device is discussed. Because of the high separation efficiencies, the rapid analysis times, and the possibility for integrating multiple chemical processes, microfluidic devices are an attractive alternative to traditional benchtop analysis. For the majority of devices, electrical potentials are utilized to actuate intricate solution flow, which lends itself to autonomous, high-throughput operation. A number of highly integrated devices have been developed that allow for the processing, concentration, and separation of analytes. In particular, devices for the processing of DNA (amplification and separation),1-3 screening for environmental toxins,4 analysis of cellular lysates,5,6 †
Geo-Centers, Inc.. Naval Research Laboratory. § Present address: Department of Chemistry, Lycoming College, Williamsport, PA 17701. (1) Khandurina, J.; McKnight, T. E.; Jacobson, S. C.; Waters, L. C.; Foote, R. S.; Ramsey, J. M. Anal. Chem. 2000, 72, 2995-3000. (2) Legally, E. T.; Emrich, C. A.; Mathies, R. A. Lab Chip 2001, 1, 102-107. (3) Ferrance, J. P.; We, Q.; Giordano, B.; Hernandez, C.; Kwok, Y.; Thiboboux, S.; Landers, J. P. Anal. Chim. Acta 2003, 500, 223-236. (4) Broyles, B. S.; Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 2003, 75, 27612767. ‡
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proteolytic digestion of proteins,7-10 and inhibition of enzymatic catalysis11 have exhibited significant potential for forensic analysis. The capability for handling real samples on microfluidic devices is a challenging hurdle which is currently restricting advancement of lab-on-a-chip devices. A number of chemical processing steps have been successfully miniaturized. They include analyte extraction,4,12-20 sample filtration,4,21,22 sample desalting,18,23,24 and analyte derivatization.25-28 Ideally, the sample processing would (5) Wheeler, A. R.; Throndset, W. R.; Whelan, R. J.; Leach, A. M.; Zare, R. N.; Liao, Y. H.; Farrell, K.; Manger, I. D.; Dariden, A. Anal. Chem. 2003, 75, 3581-3586. (6) McClain, M. A.; Culbertson, C. T.; Jacobson, S. C.; Albritton, N. L.; Sims, C. E.; Ramsey, J. M. Anal. Chem. 2003, 75, 5646-5655. (7) Gottschlich, N.; Culbertson, C. T.; McKnight, T. E.; Jacobson, S. C.; Ramsey, J. M. J. Chromatogr., B 2000, 745, 243-249. (8) Wang, C.; Oleschuk, R.; Ouchen, F.; Li, J.; Thiboult, P.; Harrison, D. J. Rapid Commun. Mass Spectrom. 2000, 14, 1377-1383. (9) Gao, J.; Xu, J.; Locascio, L. E.; Lee, C. S. Anal. Chem. 2001, 73, 26482655. (10) Peterson, D. S.; Rohr, T.; Svec, F.; Frechet, J. M. J. Anal. Chem. 2003, 75, 5328-5335. (11) Hadd, A. G.; Raymond, D. E.; Halliwell, J. W.; Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1997, 69, 3407-3412. (12) Oleschuk, R. D.; Shultz-Lockyear, L. L.; Ning, Y.; Harrison, D. J. Anal. Chem. 2000, 72, 585-590. (13) Kutter, J. P.; Jacobson, S. C.; Ramsey, J. M. J. Microcolumn Sep. 2000, 12, 93-97. (14) Cong, Y.; Davey, M. H.; Svec, F.; Frechet, J. M. J. Anal. Chem. 2001, 73, 5088-5096. (15) Jamere, A. B.; Oleschuk, R. D.; Ouchen, F.; Fajuyigbe, F.; Harrison, D. J. Electrophoresis 2002, 23, 3537-3544. (16) Ceriotti, L.; de Rooij, N. F.; Verpoorte, E. Anal. Chem. 2002, 74, 639-647. (17) Bergkuist, J.; Ekstrom, S.; Wallman, L.; Lo¨fgren, M.; Marko-Vargas, G.; Nilsson, J.; Laurell, T. Proteomics 2002, 2, 422-429. (18) Paegel, B. M.; Yeung, S. H. I.; Mathies, R. A. Anal. Chem. 2002, 74, 50925098. (19) Russom, A.; Tooke, N.; Andersson, H.; Stemme, G. J. Chromatogr., A 2002, 1014, 37-45. (20) Ro, K. W.; Cheng, W.-J.; Kim, H.; Koo, Y.-M.; Hahn, J. N. Electrophoresis 2003, 24, 3253-3259. (21) He, B.; Tan, L.; Regnier, F. Anal. Chem. 1999, 71, 1464-1468. (22) Andersson, H.; van der Wijngaart, W.; Enoksson, P.; Stemme, G. Sens. Actuators, B 2000, 67, 203-208. (23) Lion, N.; Gabry, V.; Jensen, H.; Rossier, J. S.; Girault, H. Electrophoresis 2002, 23, 3583-3588. (24) Song, S.; Singh, A. K.; Shepodd, T. J.; Kirby, B. J. Anal. Chem. 2004, 76, 2367-2373. (25) Jacobson, S. C.; Koutny, L. B.; Hergenro ¨der, R.; Moore, A. W.; Ramsey, J. M. Anal. Chem. 1994, 66, 3472-3476. (26) Jacobson, S. C.; Hergenro ¨der, R.; Moore, A. W.; Ramsey, J. M. Anal. Chem. 1994, 66, 4127-4132. (27) Fluri, K.; Fitzpatrick, G.; Chiem, N.; Harrison, D. J. Anal. Chem. 1996, 68, 4285-4290. 10.1021/ac0507789 CCC: $30.25
© 2005 American Chemical Society Published on Web 09/13/2005
be the front end of a larger device, which also couples the technique to a separation column, a mass spectrometer, or both. Because most extraction techniques allow for the removal of the analyte supporting matrix, extraction is a highly attractive candidate as a pretreatment technique for real samples. Solid-phase extraction (SPE) has been performed in microchannels by a number of researchers.4,12-20 Two important requirements of SPE are that the solid phase must reversibly retain the analyte, and it must possess sufficient surface area for the adsorption of large amounts of analyte. Small-diameter silica beads derivatized with a hydrophobic silane, for example, octadecyl (C18) liquid chromatography packings, have been commonly utilized for SPE. The real challenge has been the fabrication of microcolumns for extraction. Normally, beads have been mechanically held in place by microfabricated pillars, posts, or constrictions in the channel. Both silicon,17,22 which can be intricately fabricated through deep reactive ion etching, and polymeric materials,20 which can be molded, have been used to produce channels containing a series of posts or pillars that are closely spaced (1.5 tons) for 3 h at ∼100° C prior to final bonding at 550° C. Glass tubing, which acted as reservoirs, was affixed at the access holes using a thermally setting epoxy (353ND-T; Epoxy Technology). The layout of the device with labeling of the reservoirs is shown in Figure 1. Fabrication of Methacrylate Frits. Prior to polymerization, the chip was sequentially rinsed for 10 min with 1 M NaOH, deionized water, and methanol. After drying the microchannels, a solution consisting of 0.5% (v/v) 3-(trimethoxysilyl)propyl acrylate (covalent adhesion promoter) in toluene was loaded into the microchannels via capillary action, allowed to react for 30 min, and subsequently flushed from the channel with toluene. To prepare for the photopolymerization, two pieces of black electrical tape were used to produce a crude slit (∼200 µm wide) on the bottom of the device, which exposed the desired portion of the channel. With the exception of the exposed slit width, the bottom of the device was then completely covered with aluminum tape. At this point, a polymerization solution was prepared by combining 1.42 g glycidyl methacrylate (monomer), 0.96 g of ethylene Analytical Chemistry, Vol. 77, No. 20, October 15, 2005
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Table 1. Applied Voltages Utilized for Controlling the Extraction and Subsequent Separationa applied voltages (kV vs ground)
elution extract extract recover injection
sample
elution
buffer
elution waste
waste
5.5 6.5 6.5 5.5
6.5 5 5 6.5
3 1.5 3 3
6.5 1.8 6.5 4.5
0 0 0 0
a Details about the individual experimental steps and their order are described in the text.
Figure 1. Schematic of the microfluidic layout for the solid-phase extraction/separation device: (A) layout of the entire device and (B) expanded view of the extraction region of the device. The dotted line signifies the direction of fluid flow during extraction; the solid line signifies flow during elution/injection. The narrow channels are ∼55 µm wide; the column chamber is ∼210 µm wide. All channels are ∼15 µm deep.
dimethacrylate (cross-linker), 0.024 g of 2,2′-azobisisobutyronitrile (initiator), 0.005 g of 2-acrylamido-2-methyl-1-propane sulfonic acid (electroosmotic flow promoter), and 3.6 g of porogenic solvent (50% w/w ethanol, 50% methanol). Before preparing the polymerization mixture, each of the methacrylates was equilibrated with activated aluminum oxide to remove inhibitor. With vacuum applied to the waste (also called separation waste) reservoir, the device was filled by pipetting 10 µL of the polymerization mixture into each reservoir. Vacuum application continued until all air was removed from the channel. The reservoirs were evacuated of all solution and sealed with Parafilm, after which the device was covered with aluminum tape. The microfluidic device was placed on a cooled block of aluminum and exposed to ultraviolet radiation for 45 min using the same lamp as for photolithography. Following polymerization, the device was sequentially flushed with toluene, acetonitrile, and deionized water for 10 min. Preparation of the Solid-Phase Microcolumns. The microfluidic device was filled with acetonitrile by applying negative pressure to the waste reservoir until the frit was wetted (i.e., all air was removed). At this point, the acetonitrile in the sample and elution reservoirs was removed and replaced with a 3 mg/mL slurry of octadecyl derivatized silica beads (3 µm Luna; Phenomenex). To pack the column, negative pressure was applied to the sample waste (also called elution waste) reservoir for 10 s. The column was rinsed with acetonitrile and water for 10 min each. The beads were “locked” in place by flushing with 40 mM Tris/ HCl buffer (pH 8.8). This “solvent lock” phenomenon is similar to that described previously.12,15 Prior to analysis, the columns were conditioned with 0.01 N HCl, followed by equilibration with 6666 Analytical Chemistry, Vol. 77, No. 20, October 15, 2005
running buffer (40 mM Tris/HCl). Although care was taken to avoid introduction of air into the extraction channel, bubbles would occasionally become trapped in the SPE column and require flushing with toluene for removal. Because the sample (40 mM Tris/HCl) and elution (40 mM Tris/HCl, 80% (v/v) 2-propanol) buffers are of a different composition from the separation buffer (40 mM Tris/HCl, 20 mM sodium dodecyl sulfate), care was taken to load appropriate buffers into the appropriate channels. Briefly, the device had been filled initially with the 40 mM Tris/HCl. Just prior to analysis, the buffer, sample waste, and waste reservoirs were filled with the separation buffer (40 mM Tris/HCl, 20 mM SDS), and vacuum was applied at the waste reservoir for 5 min. The buffers in the sample and elution reservoirs were replaced with the sample (40 mM Tris/HCl, analyte) and elution buffer (40 mM Tris/HCl, 80% 2-propanol) while the remaining reservoirs were replenished with the separation buffer. Microfluidic Analysis. Analysis was driven using an array of six high-voltage supplies (2866A; Bertan), which was controlled by personal computer via a Labview program and a controller card (PCI-MIO-16XE; National Instruments). Through control of the voltages applied to each of the reservoirs, fluid continued to flow across the SPE column and at any time permitted injection of a plug of analyte onto the separation channel. The voltages required to control each of the processes are contained in Table 1. To initiate a typical experiment, elution buffer was first directed across the column (to remove any adsorbed analyte prior to analysis) through application of voltages identical to those used for the elution step. By varying the voltages at the sample and elution reservoir, either sample (extraction) or elution buffer (elution) could be directed toward the SPE column. The voltages at the sample waste and buffer reservoirs were lowered during extraction to maximize the loading of the analyte onto the column head. For the last 10 s of the extraction, the voltages were reinstated to allow for reestablishment of the gated injection valve at the expense of the flow rate through the SPE column (extraction recovery). Detection. Laser-induced fluorescence was utilized for detection of the fluorescent analytes during extraction and separations in this study. For the majority of the experiments described, the desired line of an air-cooled Ar+ (ILT5000; Ion Laser Technology) or an Ar+-Kr+ laser (35-KAP-431-208; Melles Griot) was focused into the microchannel via a 100-mm lens, while the resulting fluorescence was collected by a 20× microscope objective (numerical aperture ) 0.4). Detection was performed 5 cm below the injection cross. Analyte fluorescence was filtered spatially (1mm pinhole) and spectrally (Omega) prior to detection with a
photomultiplier tube (H7732-10; Hamamatsu). For the rhodamine dyes, 514-nm excitation was used, and the fluorescence was filtered using a 580-nm ((10) band-pass filter. For coumarin, 457nm excitation was used and filtered using a 500-nm ((10) bandpass filter. The resulting PMT signal was amplified and recorded via a second controller card (PCI-6713; National Instruments) and the same Labview routine as mentioned above. RESULTS AND DISCUSSION Porous photopolymerized acrylate and methacrylate polymers have been used extensively on microfluidic platforms as stationary phases,28,29 pressurized flow actuators,30,31 and as sorbents for SPE.14 Interestingly, plugs of these polymers have also been used as frits for retaining beads in conventional capillary electrochromatography.32 This work demonstrates the first utilization of photopolymerized frits for retaining beads in microfluidic channels for solid-phase extraction. Although intricate fabrication of methacrylate polymers has been previously performed, fabrication of the small frits necessary for microfluidic frits was challenging. Control of a number of factors was critical to success in the photopolymerization of these methacrylate monomers. The first factor was purity of the monomers. To prevent unintended polymerization of the monomer, an inhibitor was included by the supplier. Exposure to a basic alumina powder effectively removed the inhibitor and allowed for more efficient polymerization.33,34 The second factor was the need for efficient cooling of the fluidic device during exposure. Cooling was necessary to eliminate undesirable polymerization due to activation of initiators by heat as well as UV exposure. An aluminum block cooled in an ice bath was utilized both as a heat sink and for supporting the device during exposure. The third and most important factor was the elimination of localized fluid flow in the microchannels due to siphoning effects. When fluid motion occurs, reactive intermediates are swept away from the exposed region of the channel, causing sporadic polymerization. This results in irregular, elongated masses of polymer that are not appropriate for holding beads (Figure 2A). When siphoning was minimized through removal of excess fluid from the reservoirs, clean plugs of polymer with an length of ∼150 µm were produced (Figure 2B). According to previously published polymerization procedures,33 a pore size of 5 µm was expected, but no porosity measurement was attempted. The pore size was larger than the beads utilized for extraction (3 µm), but the random nature of the pores prevented the beads from passing through. Once the frit was in place, a simple procedure was used to pack the solid-phase extraction column. Pulling a slurry of beads in acetonitrile through the frit produced a well packed column in a short period of time (10 s). Figure 3A shows a column prepared using the methods described in the Experimental Section. The beads pack evenly and, in most cases, without voids. More (29) Throckmorton, D. J.; Shepodd, T. J.; Singh, A. K. Anal. Chem. 2002, 74, 784-789. (30) Kirby, B. J.; Shepodd, T. J.; Hasselbrink, E. F. J. Chromatogr., A 2002, 979, 147-154. (31) Reichmuth, D. S.; Shepodd, T. J.; Kirby, B. J. Anal. Chem. 2004, 76, 50635068. (32) Chen, J.-R.; Dulay, M. T.; Zare, R. N.; Svec, F.; Peters, E. Anal. Chem. 2000, 72, 1224-1227. (33) Yu, C.; Xu, M.; Svec, F.; Frechet, J. M. J. J. Polym. Sci. 2002, 40, 755-769. (34) Tan, A.; Benetton, S.; Henion, J. D. Anal. Chem. 2003, 75, 5504-5511.
Figure 2. Fabrication of photopolymerized frits: (A) sporadic polymerization due to siphoning and other factors (see text) and (B) successful fabrication of a usable frit.
importantly, the beads could be locked in place by flushing the column with aqueous buffer solutions. The aggregation of the beads, which is presumably due to the hydrophobic interactions, was strong enough that removal of the column was difficult. Once prepared, the same solid-phase extraction microcolumn was utilized for several weeks at a time. The initial intent had been to hold the beads in place with a second polymeric frit, but the forces holding the column together were strong enough to withstand the electrophoretic forces on the weakly negatively charged beads in the presence of a strong electric field. Figure 3B shows a fluorescence image of the microcolumn shown in part A using the same imaging power and after electrophoretic loading with rhodamine B (10 µM) for 10 min. Figure 3B illustrates the strong association of the beads even in an applied electric field. This behavior is contrary to previously published SPE studies in which the beads were held in place by two mechanical barriers12,15 to prevent the beads from electrophoresing toward the anode. Due to the dimensions of the channels utilized in the microfluidic devices, small-diameter beads were necessary. Although small-diameter beads possess high surface areas and lead to efficient chromatographic separations, columns of these beads can generate high back pressures in pressure-driven systems. In electrically controlled systems, back pressures can also result from end capping. Because end capping, the process of reducing silanol concentration, effectively reduces the ζ potential, the beads have less ability to support electroosmotic flow than the open channel walls. Nonuniform ζ potentials have been shown to generate pressures in capillary electrophoresis. Initial work was performed Analytical Chemistry, Vol. 77, No. 20, October 15, 2005
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Figure 4. Quantitative solid-phase extraction of rhodamine B on a microfluidic device. Three experiments are shown with varying extraction times (30, 60, 90 s). The open circles represent the average of three successive extractions with errors in intensity and elution time indicated with bars. The straight line is a linear fit of the average data (y ) 1.94x - 32.4; r ) 0.99). The band marked with an asterisk (*) is carryover from a previous run. Figure 3. Loading of columns with beads and their use: (A) microfrit retaining 3-µm-diameter beads and (B) fluorescence image of the microcolumn shown in part A using the same imaging power and after electrophoretic loading with rhodamine B (10 µM) for 10 min.
in narrow, uniform width channels that gave high back pressure during both column packing and extraction. To minimize the back pressure, the channel design was altered to provide a widened chamber for containing the column (Figure 1B). Although the expansion ratio utilized (55 µm/210 µm) was not optimized, the back pressures were minimized, and the devices functioned in a fashion similar to devices without columns. The column performance for solid-phase extraction was quite good. As noted earlier, the columns were stable at high electric field strengths, at differing organic solvent concentrations, and following repeated use. After 10 min of loading a relatively high concentration of fluorescent dye (Figure 3B; rhodamine B; 10 µM), less than 10% of the column capacity had been consumed. To determine the capacity, an attempt was made to measure analyte breakthrough. After loading 1 µM rhodamine B onto the column for 45 min (16 pmol), no apparent breakthrough was observed. For comparison, previously published capacities were 125 fmol for derivatized open channels,4 190 pmol for monolithic polymers,14 and 81 fmol for packed-bead columns.15 It is not clear why the capacity of the columns used for this study is so much larger than those of other bead-based microextractions.15 The monolithic polymers utilized in ref 14 were much longer, however, which explains their large capacities. Figure 4 shows the effect of analyte loading time on the observed fluorescence in the form of both the actual data and the corresponding linear regression. As expected, the signal tracks linearly with loading time. For the 100 nM rhodamine B solutions utilized, the signal reaches the instrumental maximum after less 6668 Analytical Chemistry, Vol. 77, No. 20, October 15, 2005
than 120 s of extraction. A preconcentration factor (PF) can be defined as the ratio of the solution volume passed through the column during extraction to the volume used for eluting the analyte.12,15 For the extractions in Figure 4, the preconcentration factors ranged from 70 for the 30-s extraction to 220 for the 90-s extraction. The calculation relies on measured flow rates (νextraction ) 6.2 nL/s; νelution ) 1.1 nL/s) and an eluted bandwidth of 2.35 s (average full width at half-maximum of a Gaussian fit). With high preconcentration factors, the extraction of trace analytes can be performed. Figure 5 illustrates the extraction of a 100 pM solution of rhodamine B with an extraction time of 300 s. The signal-tonoise ratio (S/N) observed was in excess of 5000. Assuming a linear dependence of S/N ratio on concentration, a theoretical detection limit of 60 fM was calculated. Columns prepared using methacrylate frits were reproducible in both dimension and performance. When prepared using the procedures outlined in the Experimental Section, columns of similar length could readily be prepared. For a bead-loading time of 10 s, columns of similar length were repeatedly produced (not shown). The error bars in Figure 4 illustrate the reproducibility of the extractions. The relative error in elution times was within 3% for the data presented, whereas the relative error in signal intensity was less than 15%. For trace analysis in environmental, forensic, and antiterrorist analytical applications, it would be valuable to autonomously integrate sample processing with chemical analysis. Although this has been previously achieved,4 extraction was performed on the same column that supported the separation. It can be disadvantageous to utilize the same column for both extraction and separation, mainly because as the column is loaded, the effective separation distance is reduced, and the separation efficiency is
Figure 5. Trace analysis of 100 pM rhodamine B. The analyte solution was extracted for 300 s.
diminished.4,35,36 It is anticipated that this would be a problem when a significant fraction of the column capacity is consumed by the on-column extraction, that is, at long extraction times. Additionally, because the beads were not held in place mechanically, microscopic shifting of the column packing was occasionally observed. Although this rarely affected the ability to perform SPE, any motion of the column would be highly deleterious to separation efficiencies and run-to-run reproducibility. This led us to investigate the application of a second channel for performing MEKC separations, as opposed to simply utilizing the packed column for electrochromatography. A gated injection valve was used to interface the extraction and separation columns. Gated injection is a variable volume injection method that allows the user to select a reproducible plug length. As the concentrated band of analyte passed through the valve, an injection was made onto the separation column. Therefore, it was necessary that the elution time be known and reproducible. The elution time was first measured immediately prior to the injection valve. As the nonaqueous eluting solvent evaporated from the eluent reservoir, the elution time was observed to shift to longer times. This is expected as the partition coefficients change with changing organic elutant concentration. To ensure reproducibility, the elution buffer was refreshed immediately prior to performing each experiment. Because of the success of reversed-phase separation techniques in the analysis of toxic analytes of concern (environmental/ polyaromatic hydrocarbons; forensic/explosives; antiterrorism/ warfare agents), micellar electrokinetic chromatography (MEKC) was chosen as the separation method. Previous efforts by this group have demonstrated the application of C18-coated beads as an effective solid-phase material for the extraction and preconcentration of explosives from seawater. A series of neutral, (35) Seifer, R. M.; Kraak, J. C.; Kok, W. T.; Poppe, H. J. Chromatogr., A 1998, 808, 71-77. (36) Liu, Z.; Otsuka, K.; Terabe, S. Electrophoresis 2001, 22, 3791-3797.
Figure 6. Separation of preconcentrated dyes. Coumarin 314 and 334 were used as analytes and extracted for 60 s prior to separation on the same device. The analytes were separated via MEKC. The bands marked with an asterisk (*) are carryover from a previous run.
fluorescent dyes were chosen to mimic the extraction behavior of organic contaminants, such as explosives; permit direct fluorescence microscope imaging of the microchip’s performance; and, finally, allow for comparison with previous microchip extractions, many of which have utilized neutral dyes. Autonomous extraction and separation of coumarin 314 and 334 is shown in Figure 6. The dyes were present at 100 nM and were hydrophobically extracted for 60 s. The performance of the device is quite good with high separation efficiencies observed for the separation. The observed separation efficiency for coumarin 334 (the first band) was 32 000 plates for a plate height of 1.6 µm (640 000 plates/m). For coumarin 314, 38 000 plates was observed for a plate height of 1.3 µm (760 000 plates/m). These efficiencies are similar to those observed previously for microfluidic MEKC separations.37,38 It is apparent that the dye bands are not identical in intensity. This can be attributed to differing partition coefficients for the coumarin dyes, which could lead to the observed differences in elution times. Even a slight difference in elution timing between the analytes can cause shifts in the relative intensity of analytes, and work is continuing to minimize this variability. It is also important to note that the signal-to-noise ratios for the coumarin dyes are not as high as those observed for the rhodamine dyes. This is mainly due to the fact that the violet laser line (457 nm) used to excite the coumarins is not as intense as the green line (514.5 nm) utilized for rhodamine. The autonomous nature of this device is a significant improvement over macroscale solid-phase extraction techniques. Previous work by this group was directed toward rapid, semiautomated solid-phase extraction techniques.39 By integrating chemical (37) Kutter, J. P.; Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1997, 69, 51655171. (38) Ramsey, J. D.; Jacobson, S. C.; Culbertson, C. T.; Ramsey, J. M. Anal. Chem. 2003, 75, 3758-3764. (39) Smith, M.; Collins, G. E.; Wang, J. J. Chromatogr., A 2003, 991, 159-167.
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analysis directly onto the same device, the current work represents, at minimum, a 3-fold improvement in time over previous efforts. As Figure 6 illustrates, the entire pretreatment and analysis takes under 4 min. Additionally, the time savings are greater than the simple sum of extraction and analysis times due to the elimination of user intervention. Devices of this type could conceivably be utilized in field-based forensic or combat situations, in which the operator would not be a traditional research chemist or for which minimal operator intervention is desired. CONCLUSION The first demonstration of a completely automated microdevice which integrates micro-SPE sampling, with on-chip elution, sample injection, and separation of neutral compounds by micellar electrokinetic chromatography (MEKC) was shown. Trace analysis in the fields of environmental analysis, forensic science, and terrorism prevention will benefit significantly from the integration of on-chip preconcentration elements with miniaturized, automated injection and separation capabilities. Nonaqueous capillary electrophoresis separations will have a direct and immediate impact
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on the on-chip SPE results discussed here, due to the resulting compatibility of the eluent and separation buffer phases. The challenge now is to optimize the performance of these devices, and widen their applicability toward real samples. ACKNOWLEDGMENT The authors thank Dr. Qin Lu, Dr. Braden Giordano, and Dr. Christopher Tipple for useful discussions during preparation of this manuscript. Funding support was provided by the Office of Naval Research (ONR) and the Memorial Institute for the Prevention of Terrorism (MIPT). Points of view in this document are those of the authors and do not necessarily represent the official position of the U.S. Department of Homeland Security or MIPT.
Received for review May 5, 2005. Accepted August 12, 2005. AC0507789
