Concentration of DNA in a Flowing Stream for High-Sensitivity

Maria Rowena N. Monton , Shigeru Terabe. Journal of Chromatography B ... E. J. Walsh , C. King , R. Grimes , A. Gonzalez. Biomedical Microdevices 2006...
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Anal. Chem. 2003, 75, 4467-4474

Concentration of DNA in a Flowing Stream for High-Sensitivity Capillary Electrophoresis Sang-Ryoul Park† and Harold Swerdlow*,‡

Department of Human Genetics, University of Utah, 20 S. 2030 E., Room 308, Salt Lake City, Utah 84112-9454

A novel sample pretreatment device is described, and its application to the concentration and purification of crude DNA samples in a flowing stream for subsequent capillary electrophoresis is demonstrated. The device consists of two gap junctions, each covered with a conductive membrane material and built upon a flow channel made of PEEK tubing. Upon the application of an electric field between the junctions, the negatively charged DNA fragments can resist the hydrodynamic flow stream and are trapped between the junctions. DNA fragments dissolved in microliter volumes are captured in a nanoliter-sized band by simply pushing the sample solution through the device. Depending on their electrophoretic mobility, other interfering materials in a crude sample can be removed from the trapped DNA fragments by washing. The selective permeability of the membrane to small ions allows efficient desalting. The concentrated and purified DNA fragments are released by simply turning off or reversing the electric field. Recovery is up to 95%. Performance of the device was evaluated using crude products of fluorescent dye-primer cycle-sequencing reactions. Compared to these crude reaction products, samples purified in the capture device and subsequently collected showed dramatically enhanced signal and resolution when run on a conventional capillary-electrophoresis instrument. Furthermore, the device could be connected in-line to a capillary system for direct injection. The device has great potential for enabling lab-on-a-chip systems to be used with real-world samples. The high resolving power of DNA electrophoresis has been largely responsible for many recent spectacular advances in genotyping, polymerase chain reaction (PCR) analysis, restriction fragment digest analysis, mutation screening, and DNA sequencing.1,2 Although conventional slab-gel electrophoresis is still widely used for DNA sequencing, several completed genomes attest to the ease of operation, speed of separation, lower reagent costs, † Current address: Div. of Chemical Metrology, Analytical Laboratory for Biological/Clinical Applications, Korea Research Institute of Standards and Science, P. O. Box 102, Yuseong, Daejeon, 305-600, S. Korea. Fax: +82 42 868 5042. E-mail: [email protected]. ‡ Current address: Solexa Ltd., Chesterford Research Park, Little Chesterford, Essex, CB10 1XL, U.K. Fax: +44 1799 532 306. E-mail: harold.swerdlow@ solexa.com. (1) Cohen, A. S.; Najarian, D. R.; Paulus, A.; Guttman, A.; Smith, J. A.; Karger, B. L. Proc. Natl. Acad. Sci. U.S.A. 1988, 85, 9660-9663. (2) Smith, L. M. Curr. Opin. Biotechnol. 1993, 4, 37-40.

10.1021/ac034209h CCC: $25.00 Published on Web 07/22/2003

© 2003 American Chemical Society

and superior resolving power of capillary-electrophoresis (CE) techniques.3-5 Developments for using replaceable linear-polymerbuffer systems rather than conventional cross-linked gels in the capillary made it possible to accomplish up to 150 runs without replacing the separation capillary and furthermore eliminated gel pouring, one of the most time-consuming and labor-intensive steps in slab-gel electrophoresis.6-9 Throughput enhancement is achieved through the use of multiple separation capillaries (96 or 384) in a single CE instrument and multiple unattended runs.10-14 Despite these advantages, progress in overall throughput and cost for DNA sequencing projects has been limited to a great extent by the paucity of methods for efficiently preparing and purifying small-volume samples. CE performance becomes severely degraded when products of biochemical reactions (e.g., DNA sequencing or PCR) are loaded without appropriate sample pretreatment.15-17 Furthermore, unlike slab-gel electrophoresis in which virtually the entire sample in the well enters the gel, loading efficiency in CE is inversely proportional to the salt (and buffer) concentration of the sample. This is explained, at least in part, by exclusion of DNA fragments from the gel by faster migrating salt ions in a typical electrokinetic loading.18 Therefore, purification of reaction products is routinely performed and is essential for high-sensitivity CE. Precipitation of DNA fragments with cold ethanol, followed by washing, is usually adequate to eliminate salt.4 (3) Luckey, J. A.; Drossman, H.; Kostichka, A. J.; Mead, D. A.; D’Cunha, J.; Norris, T. B.; Smith, L. M. Nucleic Acids Res. 1990, 18, 4417-4421. (4) Swerdlow, H.; Gesteland, R. Nucleic Acids Res. 1990, 18, 1415-1419. (5) Cohen, A. S.; Najarian, D. R.; Karger, B. L. J. Chromatogr. 1990, 516, 4960. (6) Pariat, Y. F.; Berka, J.; Heiger, D. N.; Schmitt, T.; Vilenchik, M.; Cohen, A. S.; Foret, F.; Karger, B. L. J. Chromatogr., A 1993, 652, 57-66. (7) Fung, E. N.; Yeung, E. S. Anal. Chem. 1995, 67, 1913-1919. (8) Menchen, S.; Johnson, B.; Winnik, M. A.; Xu, B. Electrophoresis 1996, 17, 1451-1459. (9) Best, N.; Arriaga, E.; Chen, D. Y.; Dovichi, N. J. Anal. Chem. 1994, 66, 4063-4067. (10) Ueno, K.; Yeung, E. S. Anal. Chem. 1994, 66, 1424-1431. (11) Zhang, J.; Voss, K. O.; Shaw, D. F.; Roos, K. P.; Lewis, D. F.; Yan, J.; Jiang, R.; Ren, H.; Hou, J. Y.; Fang, Y.; Puyang, X.; Ahmadzadeh, H.; Dovichi, N. J. Nucleic Acids Res. 1999, 27, e36. (12) Nay, L. M.; Sinclair, R.; Swerdlow, H. Proc. SPIE 1997, 2985, 19-24. (13) Huang, X. C.; Quesada, M. A.; Mathies, R. A. Anal. Chem. 1992, 64, 21492154. (14) Bashkin, J. S.; Bartosiewicz, M.; Roach, D.; Leong, J.; Barker, D.; Johnston, R. J. Capillary Electrophor. 1996, 3, 61-68. (15) Swerdlow, H.; Dew-Jager, K. E.; Brady, K.; Grey, R.; Dovichi, N. J.; Gesteland, R. Electrophoresis 1992, 13, 475-483. (16) Ruiz-Martinez, M. C.; Salas-Solano, O.; Carrilho, E.; Kotler, L.; Karger, B. L. Anal. Chem. 1998, 70, 1516-1527. (17) Salas-Solano, O.; Ruiz-Martinez, M. C.; Carrilho, E.; Kotler, L.; Karger, B. L. Anal. Chem. 1998, 70, 1528-1535. (18) Xiong, Y.; Park, S.-R.; Swerdlow, H. Anal. Chem. 1998, 70, 3605-3611.

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Further, because the adverse effect of injection of template DNA on sequencing performance in both slab gels and capillaries is well documented,15,19,20 the DNA fragments can be resuspended in, for example, a template suppression reagent (TSR), as supplied by Applied Biosystems.8 Alternatively, purification of DNA sequencing reaction products from template DNA and small ions using ultrafiltration, spin-columns, or magnetic beads has been reported to be very effective, resulting in a 10-50-fold increase in DNA injection.16,17,21 Nevertheless, these DNA sample treatments are still laborious and time-consuming and can represent a bottleneck in the complete DNA sequencing operation. Various strategies have been suggested to improve the basic methods for sample preparation and purification in CE. For example, we have developed a sample-loading technique, based on base-stacking of DNA, that eliminates the need for intensive sample pretreatment.18 Numerous groups have also reported fluidic integration on both capillary and chip-based systems. Automation of the biochemical reaction, purification, sample injection, separation, and detection steps are achieved by coupling these components together fluidically. Typically, these systems aim to concentrate samples between the subsystems. The first 2-D systems based upon CE as the second dimension were described by Jorgenson’s group.22 A capillary-based on-line system for concentration and desalting of proteins by microvolume electrodialysis was described by Wu et al.,23 and Waldron et al. have described a fluidic CE system that can purify and concentrate proteins based upon inline solid-phase extraction.24 Mathies, Northrup, and colleagues have created coupled PCR and electrophoretic separation devices on chips.25 Mathies has also created a method for in-line purification of DNA based upon hybridization to complementary DNA bound to a gel matrix in a chip format.26 Purification and concentration is complete in only 2 min. Ramsey’s group has reported an elegant in-situ cleanup technique for chips based upon a porous polysilicon membrane-based salt bridge.27 Meldrum has created an automated capillary-based reaction system that, although not in-line, is nonetheless a sophisticated, robust instrument.28 We have pioneered the use of direct in-line loading of PCR and sequencing reactions onto a CE system via fluidic integration.29 This automated system consists of an air thermal cycler, an HPLC pump, a gel-filtration column, two switching valves, a flow splitter, and a CE subsystem. Briefly, the PCR or DNA cyclesequencing reaction is performed in the loop of an injection valve situated inside the air thermal cycler. After the reaction, the (19) Swerdlow, H.; Dew-Jager, K.; Gesteland, R. F. BioTechniques 1994, 16, 684693. (20) Tong, X.; Smith, L. M. DNA Sequence 1993, 4, 151-162. (21) Devaney, J. M.; Marino, M. A.; Williams, P. E.; Weaver, K. R.; Del Rio, S. A.; Turner, K. A.; Belgrader, P. Appl. Theor. Electrophor. 1996, 6, 11-14. (22) Larmann, J. P., Jr.; Lemmo, A. V.; Moore, A. W., Jr.; Jorgenson, J. W. Electrophoresis 1993, 14, 439-447. (23) Wu, X.-Z.; Hosaka, A.; Hobo, T. Anal. Chem. 1998, 70, 2081-2084. (24) Bonneil, E.; Waldron, K. C. J. Capillary Electrophor. 1999, 6, 61-73. (25) Woolley, A. T.; Hadley, D.; Landre, P.; deMello, A. J.; Mathies, R. A.; Northrup, M. A. Anal. Chem. 1996, 68, 4081-4086. (26) Paegel, B. M.; Yeung, S. H.; Mathies, R. A. Anal. Chem. 2002, 74, 50925098. (27) Khandurina, J.; McKnight, T. E.; Jacobson, S. C.; Waters, L. C.; Foote, R. S.; Ramsey, J. M. Anal. Chem. 2000, 72, 2995-3000. (28) Meldrum, D. R. IEEE. Eng. Med. Biol. 1995, July/August, 443-448. (29) Swerdlow, H.; Jones, B. J.; Wittwer, C. T. Anal. Chem. 1997, 69, 848-855.

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product is pumped onto a gel-filtration column in which the DNA sample is purified and desalted. As it elutes from the column, the purified band of DNA is directed to a flow splitter that both connects the fluidic system to the CE system and allows a portion of the sample to be injected onto the CE column electrokinetically. This fluidically integrated system is fast, simple to operate, efficient, provides a high degree of reproducibility, and isolates the capillary from the negative effects of dust, contaminants, and crude reaction products. Several improvements could, however, be imagined in this system. To begin with, a number of expensive elements (e.g., multiport switching valves and the size-exclusion column) should be eliminated to make the system more affordable. This becomes ever more critical as the number of samples analyzed in parallel increases. Second, the mismatch between the volume of the reaction (microliters) and the volume of sample loaded onto the capillary (nanoliters) should be improved in order to achieve further reduced costs. Last, the entire system would benefit from being made compatible with micromachining techniques as used in “lab-on-a-chip” devices.25,32,33 Yeung’s group has also reported coupled capillary systems based upon similar designs.30,31 To address the shortfalls of our previous system, we have developed a new approach to sample pretreatment. This paradigm utilizes trapping (capture) of DNA fragments at a specific location between the reaction and separation subsystems in an integrated, flowing system. Capture of the negatively charged products of, for example, PCR or DNA sequencing reactions is accomplished by applying an electric field over a short region within a flow channel that is strong enough for the products to resist the hydrodynamic fluid flow forces. Creation of a capture device in the tubing or capillary flow channel is quite simple and inexpensive. The associated instrumentation uses relatively inexpensive components that can be shared between multiple flow channels. Control of this DNA capture system is straightforward. Furthermore, the fluidic system can be miniaturized to reduce reagent consumption, being compatible with integration on micromachined devices. In addition to concentration of DNA fragments in the capture region, significant desalting has been shown, a direct result of the size-specific permeability of the conducting membrane used to construct the trap. Using CE separation of the crude products of a thermally cycled DNA-sequencing reaction, we have investigated the characteristics of this DNA capture system. A counter flow focusing (CFF) system that could in principle be adapted to purification for CE was also reported by Tian et al.34 In this device, a wedge-shaped structure made of ionexchange resin was used in conjunction with an electric field to concentrate and focus samples by ∼50- or 100-fold. EXPERIMENTAL SECTION Preparation of In-Line DNA-Capture Devices. Forming the capture region involves creating two junction zones or gaps (one (30) Tan, H.; Yeung, E. S. Anal. Chem. 1997, 69, 664-674. (31) Hashimoto, M.; He, Y.; Yeung, E. S. Nucleic Acids Res. 2003, 31, e41. (32) Waters, L. C.; Jacobson, S. C.; Kroutchinina, N.; Khandurina, J.; Foote, R. S.; Ramsey, J. M. Anal. Chem. 1998, 70, 158-162. (33) Harrison, D. J.; Fluri, K.; Seiler, K.; Fan, Z.; Effenhauser, C. S.; Manz, A. Science 1993, 261, 895-897. (34) Tian, Z. W.; Lin, H. S.; Chen, D. Y.; Zhou, Y. L.; Mao, B. W.; Chen, H. Proc. Electrochem. Soc. 1997, 97-19, 319-323. See also http://home.att.net/ ∼bob.stevenson/99alnapce98.html

Figure 1. Schematic illustration of the major fluidic components of the DNA-capture device. Associated pumps, power supplies, etc., have been omitted for clarity.

for the upstream anode and one for the downstream cathode; see Figure 1) in the walls of an otherwise continuous flow channel. The gaps are covered with an ion-permeable membrane material to allow current to pass from the electrodes located in the buffer chambers through the gaps and into the channel, creating an electric field in the region between the gap junctions. These junctions were made by covering two small openings in a piece of PEEK tubing (Catalogue no. PM-1840; Upchurch Scientific, Oak Harbor, WA) with Nafion tubing, which swells in ethanol and shrinks upon drying (Catalogue no. TT-020; Perma Pure, Toms River, NJ). The PEEK tubing was 125 µm i.d., 635 µm o.d., and 6 cm in length; the dimensions of the two pieces of Nafion tubing were 330 µm i.d., 510 µm o.d., and 0.5 cm in length when in a dried state. The upstream opening was prepared by cutting a halfthickness notch in the PEEK tubing with a single-edged razor blade. The downstream opening was created by completely cutting the PEEK tubing and separating the two pieces by 0.2 mm. The distance between the two openings was typically 2.6 cm. The Nafion was immersed in 95% ethanol for a few minutes in order to make it fit over the PEEK tubing. The swollen Nafion tubing was slipped over the notch or gap in the PEEK tubing. Air-drying the Nafion tubing resulted in tight seals over the openings. The device was glued into a microtube rack (Catalogue no. 36010; Midwest Scientific, Valley Park, MO) for support, and two of the wells served as electrode chambers. Platinum-wire electrodes were fitted to the two microtube rack wells, and the wells were each filled with 3 mL of 100 mM Tris/ HCl, pH 8.0. The two electrodes were connected to a high-voltage power supply (Catalogue no. MJ30N0400; Glassman High Voltage, Whitehouse Station, NJ) such that the upstream electrode was positive with respect to the downstream electrode. The ends of the PEEK tubing were connected to fused-silica capillaries (365375 µm o.d.; Polymicro Technologies, Tucson, AZ) via a short piece of Teflon tubing (300 µm i.d.; Cole-Parmer, Vernon Hills, IL). The inlet and outlet ends of the PEEK tubing were sharpened to a conical shape using 400-grit sandpaper to allow them to be more easily inserted into the Teflon fittings. DNA Capture and Recovery. Crude reaction products were loaded onto the DNA-capture device by applying air pressure to a small-volume sample vial that was connected to the device through a fused-silica capillary. The volume of sample loaded onto the device was adjusted by varying the length and the inner diameter of the interconnecting capillary. The connection of this capillary to the sample vial port was made with a micro fingertight fitting (Upchurch Scientific), which facilitated easy connection/disconnection. After sample loading, flow through the capture device was produced by a model 33 syringe pump (Harvard Apparatus, Holliston, MA), using a 1-mL gastight syringe (Hamil-

ton Corp., Reno, NV). The syringe was filled with either deionized water or between 2 and 8 M urea solution. DNA fragments were captured from the flow stream by application of an electric field across the capture device. Capture and release of DNA fragments was monitored using laser-induced fluorescence (LIF) detection of the tetramethylrhodamine-labeled DNA fragments. A 1.5-mW He-Ne laser (543.5 nm, Melles Griot, Irvine, CA) was used for excitation. Fluorescence emission was collected and focused using a 20×, 0.4 NA microscope objective (Newport Optical, Irvine, CA), passed through a 1.6-mm-diameter pinhole, a band-pass filter (Catalogue no. 580DF10; 580-nm center wavelength, 10-nm bandwidth; Omega Optical, Brattleboro, VT), and a long-pass filter (OG 570; Omega Optical). Fluorescence was detected using a photomultiplier tube (Catalogue no. R1477; Hamamatsu, Bridgewater, NJ), and the photocurrent was converted to a voltage signal that was subsequently low-pass-filtered at 5.5 Hz. The fluorescence detection window ∼5 cm downstream of the capture device was made by attaching a piece of 100-µm-i.d., 375-µm-o.d. fused-silica capillary with UV-transparent coating (Catalogue no. TSU100375; Polymicro Technologies) to the outlet of the device. Electrical conductivity of the solution eluting from the downstream end of the device was continuously monitored using a homemade conductivity meter with a miniature conductivity probe in direct contact with the solution.35 Current in the capture region (between the 2 electrodes) was also monitored using a homemade current-tovoltage converter circuit that included a circuit breaker that would shut down the high voltage power supply if an excessively large or sudden change in current were detected. Signals from the fluorescence, conductivity, and current detectors were measured with an analogue to digital converter/data acquisition board (Catalogue no. DAS8-PGA; Keithley Metrabyte, Taunton, MA) and Labtech Notebook software (Laboratory Technologies, Wilmington, MA). DNA fragments were released from the capture device by either turning off the power supply or reversing its polarity. The released DNA band was either pumped from the outlet of the device into a 0.5-mL microcentrifuge tube for subsequent analysis by CE using conventional mode electrokinetic loading or coupled directly to the CE system. CE Analysis. Conventional mode CE was performed with either an ABI 310 genetic analyzer (Applied Biosystems, Foster City, CA) or a homemade CE system using the LIF detector described above. For all CE separations, a 50-µm-i.d., 375-µm-o.d., 36-cm effective length, 47-cm length uncoated fused-silica capillary (Polymicro Technologies) was used. The capillary was preconditioned with 10% HCl and rinsed with deionized water. Separations were performed using POP-6 polymer (Applied Biosystems) as the sieving medium; the polymer was replaced after every run. Captured and collected samples were electrokinetically loaded for 10 s using 1.5 kV, and subsequently, 15 kV (319 V/cm) was applied for electrophoresis. In the direct coupling mode, the outlet of the device was aligned with the injection end of the separation capillary using a loose-fitting piece of Teflon tubing surrounding both; excess liquid flowed out of this Teflon tube to waste. During the CE sample (35) Park, S.-R.; Swerdlow, H. A miniature electrolytic conductivity probe using modified bipolar pulse conductometry for microliter-volume samples; Unpublished work, University of Utah, Salt Lake City, UT, 1998.

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injection phase, that is, immediately following release, the downstream electrode (cathode) of the capture device and the injection end (cathode) of the CE system were both disconnected from the power supply. Electrokinetic injection involved application of voltage between the upstream electrode of the capture device (now functioning as a cathode) and the anode buffer chamber of the CE system. In this manner, sample injection was performed for 2 min at 15 kV and with a flow of 0.3 µL/min through the capture device. After sample loading, separation was performed using the CE anode at ground potential, and a negative high voltage (15 kV) was applied to the CE cathode. The homemade LIF system described above was used for all direct-coupled experiments. Cycle-Sequencing Reactions and Sample Treatment. Singlecolor “G” cycle-sequencing reactions using M13mp18 template DNA, dideoxy-GTP, and TAMRA (tetramethylrhodamine) dyelabeled universal primer (Applied Biosystems) were prepared as described previously.36 The products from the equivalent of many separate reactions were pooled together and frozen in small aliquots at -20 °C without any other manipulation (“crude reaction products”). A small portion of this material was thawed for each day’s experiments. A second large sample was purified in a conventional manner by adding 1/10 volume of 3 M sodium acetate, pH 4.5, and 3 volumes of cold ethanol. After precipitation for 1 h at -20 °C, the DNA was recovered by centrifugation at 14000g for 15 min. The DNA pellet was washed in 70% ethanol, air-dried for 10 min, and resuspended in deionized formamide (“purified reaction products”). RESULTS AND DISCUSSION DNA Capture Device Fabrication. DNA capture devices could be easily and quickly prepared without any specialized equipment (Figure 1). As described in the Experimental Section, cation-permeable Nafion tubing was placed over two electrically conductive but fluid-tight junctions in a flow tube. Capture of DNA was accomplished by applying an electric field in the channel, using these junctions to connect to an external power supply. The electrophoretic force on the DNA was sufficient to counteract the flow at reasonably low flow rates and reasonably high electric fields. The upstream junction was formed by nicking the PEEK tubing, thereby exposing the inner channel, but maintaining the structural integrity of the tube. The downstream gap junction was made by cutting the tubing and separating the two halves by a few hundred micrometers. Shrinkage of the Nafion tubing upon drying from ethanol resulted in a tight glueless seal over the PEEK tubing. Subsequent immersion of the membrane-covered junctions in aqueous buffer solutions neither swelled the membranes nor loosened the joints. Degassing of newly formed joints by applying slight negative pressure to the ends of the tubing was often necessary to remove trapped bubbles and to render the current more stable. During use, there is a strong electric field in the channel near the cathodic junction. Localized power dissipation at this point was occasionally high enough to produce new bubbles. Increasing the cathodic gap helped to alleviate this problem. Additionally, the capture current monitoring circuit was employed as a bubble detector spontaneous formation of new bubbles caused a rather large dip (36) Swerdlow, H.; Dew-Jager, K. E.; Gesteland, R. BioTechniques 1993, 15, 512519.

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in current through the channel. When such a transient was detected, the run was automatically terminated, and the highvoltage power supply was turned off, avoiding damage to the device during unattended operation. When used with the bubble detection circuit, each DNA capture device typically lasted for several months. Electrical Properties of the Capture Device. The electrical conductance of Nafion membranes was sufficient for this application; channel current was not substantially reduced by incorporation of the membrane. For example, the total device current at typical applied voltages and sufficiently high flow rates, measured with and without the Nafion membrane, was 125 µA and 120 µA, respectively. At low flow rates, the capture device showed complex electrical properties. These properties can be explained by preferential transport of cations across the negatively charged Nafion membrane. Whereas the capture current is carried by both cations and anions almost equally along the fluidic channel, it is carried preferentially by cations across the Nafion membrane itself. At the cathodic junction, more cations are pumped out from the channel toward the electrode than migrate along the channel from the anodic end. At the same time, fewer anions are pumped into the channel across the membrane from the cathodic chamber than migrate away along the channel toward the anodic end. This phenomenon leads to depletion of both types of ions at the cathodic (downstream) junction, whereas by a similar argument, ions become more concentrated at the anodic junction. Macroscopic electroneutrality is maintained at both junctions. The depletion zone grows in length along the fluidic channel, unless hydrodynamic flow of fresh buffer through the channel can replace ions faster than they are lost electrokinetically. With stopped flow or a low flow rate, depletion of ions near the cathodic junction dramatically increases the overall electrical resistance through the capture device. As flow rate increases, resistance around the cathodic junction decreases, and the capture current increases. Below ∼1 µL/min, there is a positive correlation between the magnitude of the peak capture current and the hydrodynamic flow rate (Figure 2A), explained by the shortening of the effective length of the depleted zone at the cathodic junction. For flow rates greater than ∼1 µL/min, the current drops with increasing flow rate, to a constant value due only to the injected buffer, likely as a result of sweeping out from the channel of the ion concentrated zone. An alternative explanation is that, at high flow rates, the effect of joule heating is reduced by removal of heat downstream by the flow; this effect would reduce the current drawn by the device. A trace of the capture current measured during a typical run is shown in Figure 2B. Immediately after applying the voltage (transient at time 0) and starting the flow of crude reaction products (see Experimental), the capture current drops as a result of depletion of ions at the cathodic junction. The capture current then increases continuously as sample ions are captured and accumulate in the fluidic channel. After switching the flow to deionized water (∼500 s), the capture current again drops as ions in the channel are eventually swept out. It is interesting to note that the changes in capture current are not smooth, but increase and decrease stepwise. Although this feature was seen at various flow rates, the magnitude of the

Figure 3. Effects of the applied voltage on entrapment of DNA fragments. A sample of crude reaction products was applied until ∼500 s, followed by distilled H2O. Applied voltages (from 0 s) are given on the figure next to each trace at a flow rate of 0.3 µL/min. Curves were displaced along the y axis for clarity.

Figure 2. Capture device current. (A) Peak capture current depends on flow rate. The buffer consisted of 53.5 mM Tris/HCl, pH 8.0, 5 mM MgCl2, 1.25 mM NaCl, and 0.1 mM Na2EDTA. The applied voltage was 600 V. (B) Changes in the electrical current through the DNA capture device during a run using a sample of crude reaction products. Conditions were as in A; flow rate 0.3 µL/min. Injection at time 0.

current was highly dependent on flow rate, as shown in Figure 2A. Although a higher flow rate is desirable for faster processing of a given volume of sample, there was a limit to flow rate due to excessive joule heating in the channel. The capture devices described here could be operated at flow rates not exceeding 0.5 µL/min. Typically, we used a flow rate of 0.3 µL/min. Capturing DNA Fragments. DNA fragments are captured from the flowing stream by the application of an electric field that exerts a sufficiently high opposing force. Depending upon the magnitude of the electric field, the DNA can be localized at different positions in the flow channel, even being forced up against the semiporous membrane. The minimum electric field for capture should be proportional to the flow rate of the solution. For example, the voltage required to capture DNA from a flow stream of 0.3 µL/min is estimated as follows. The linear velocity of the flow is calculated to be 0.04 cm/s, since the inner diameter of the flow channel is 126 µm. The free-solution electrophoretic mobility in our system of a 100-nucleotide-long single-stranded DNA was measured to be 3.0 × 10-4 cm2/V‚s (data not shown).37 (37) Stellwagen, N. C.; Gelfi, C.; Righetti, P. G. Biopolymers 1997, 42, 687-703.

This electrophoretic mobility was assumed to be a good approximation to the electrophoretic mobility of the DNA sequencing products produced by our reactions. Since electrophoretic velocity is equal to the product of electrophoretic mobility and electric field, the required strength of the electric field to just counteract the fluid flow was calculated to be 130 V/cm, corresponding to a voltage of 340 V under the assumption of a uniform electric field (electrodes 2.6 cm apart). Figure 3 shows the results of attempts to capture DNA fragments with various applied voltages and a fixed flow rate of 0.3 µL/min. Sample was added until ∼500 s. Successful capture of DNA fragments was indicated by an early (before 500 s) drop in fluorescence signal at the outlet of the capture device. As suggested by the theoretical calculation above, complete capture was not seen until the voltage was increased above 350 V (Figure 3E). In practice, some evidence of capture was seen even at voltages below the calculated value (Figure 3C,D), although capture of DNA at these voltages was not permanent. DNA capture at a lower voltage than the calculated voltage is most likely due to localized focusing of the electric field within the ion-depleted zone around the cathodic junction, in contradiction to the uniformfield assumption. Capture at the depletion zone seems to be susceptible to perturbations in flow. For example, following the replacement of sample solution with deionized H2O at 500 s, a small release of captured DNA can be seen in Figure 3D,E. Generally, a voltage ∼150 V greater than the calculated value was satisfactory to avoid spontaneous release of captured DNA. The minimum required voltage was linearly dependent on flow rate when measured between 0.1 and 1.0 µL/min (data not shown). At very high capture voltages, DNA was effectively captured but the captured fragments could not be recovered. Recovery of DNA Fragments after Capture. With respect to recovery of the captured DNA, we considered where the DNA fragments get captured and if there was a direct interaction between the DNA fragments and the membrane itself. One assumes that if the DNA fragments are positioned somewhere along the fluidic channel proper, they can be easily swept from the capture device when the capture voltage is turned off. On the Analytical Chemistry, Vol. 75, No. 17, September 1, 2003

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Figure 4. Effect of capture time on distribution of captured DNA fragments. In A, capture voltage was not applied. In B-G, the 500-V capture voltage was turned off at B, 240 s; C, 480 s; D, 720 s; E, 960 s; F, 1200 s; G, 1800 s, as indicated by the positions of the arrows. In H, the polarity of the voltage was reversed at 1800 s (inverted arrow). The flow rate used was 0.3 µL/min. The sample of crude reaction products was replaced with distilled H2O at ∼480 s (note, there is a time delay caused by the dead volume between the sample injection subsystem and the capture device). Curves were displaced along the y axis for clarity.

other hand, if they have been driven into the upstream gap, recovery would first require release into the fluidic channel; this would be evidenced by a significantly broadened band of the recovered DNA fragments. In the extreme case, if the captured DNA fragments penetrate or become entangled in the membrane, the recovered DNA band would be extremely broad, and some material loss would be inevitable. Band profiles of the recovered DNA were observed under various capture and release conditions (Figure 4). In all cases, the sample solution was replaced with a low-conductivity washing buffer after ∼8 min. Release of DNA by turning off the capture voltage before the replacement of the sample solution resulted in the DNA’s being recovered in a sharp band (Figure 4B,C). If the captured DNA fragments were released after the replacement, the band height was decreased, and significant broadening can be seen (Figure 4E-G). This can possibly be related to the observation that the low-salt solution perturbed the electric field in the device and caused the DNA to move further upstream or into the gap. Some DNA was also retained in the capture device until a reversed polarity field was applied (e.g., see band eluting at ∼25 min in Figure 4D). This effect is likely due to some entanglement of the DNA within the matrix of the membrane. A key design issue for selecting a semipermeable membrane for the electrical junctions of the capture device was its ability to provide a barrier to DNA fragment migration. A poor barrier will result in substantial loss of DNA fragments during capture. Nafion membrane has an inherent advantage over other potential barriers not related to its pore size. Nafion is highly negatively charged as a result of the presence of sulfonyl groups in the polymer. Effective blocking of DNA molecules is expected as a result of strong charge-charge repulsion. This “charge exclusion” is consistent with the observation that DNA is still efficiently recovered, even after prolonged application of the capture voltage. 4472 Analytical Chemistry, Vol. 75, No. 17, September 1, 2003

Figure 5. Sharpening of the released DNA fragment band by inclusion of large DNA molecules in the sample. A, no added DNA; B, 10 ng/µL M13mp18 DNA; C, 20 ng/µL M13mp18 DNA; D, 16 ng/ µL retroviral clone DNA. M13mp18 DNA is single-stranded and ∼7000 nucleotides long. The retroviral clone DNA is double-stranded and ∼8100 bp long. Conditions are as described in the legend to Figure 4, except the capture voltage was turned off at ∼1200 s. The sample solution in this case consisted of 0.08 µM dye-labeled M13mp18 forward universal primer in DNA sequencing reaction buffer (Applied Biosystems). The peaks following the major peaks (just past 1800 s) were released by reversing the polarity of the electric field. Curves were displaced along the y axis for clarity.

For example, >90% of DNA fragments were recovered after capturing for longer than an hour (data not shown). However, it is also likely that partial penetration of the membrane occurs, resulting in the retention of a portion of the captured DNA fragments. The pore size of Nafion membranes is known to be in the range of 40-60 Å.38,39 Although the molecular weight of a typical DNA molecule is relatively large (∼30 000 Da for a 100bp single-stranded DNA), the diameter of a stretched singlestranded DNA under a reasonably high electric field is probably small enough to allow some entry into the pores of the Nafion membrane. However, the depth of penetration was not sufficient to prevent recovery of the fragments with just a short electric field polarity reversal. Furthermore, adding large DNA fragments into the sample solution dramatically reduced this phenomenon. As shown in Figure 5, addition of either M13mp18 template DNA or double-stranded DNA of 8100 bp completely removed the tailing effect of the captured DNA fragments. This phenomenon is likely due to the effective blockage of the Nafion pores by the excess added DNA. A further improvement in recovery performance may be achievable by optimization of the surface properties of the Nafion. The recovery of DNA could be as high as 95% under optimal conditions, as measured by integrating the eluted band relative to the signal from the sample DNA lost during capture. The volume of the loaded sample was 2.7 µL, and that of the recovered DNA band was measured to be only 0.55 µL, an overall concentration factor of ∼5. There is likely to be a large dilution on recovery due to laminar-flow band-broadening in the tubing leading up to the detector. Band volume in the device itself is expected to be (38) Mauritz, K. A.; Storey, R. F.; Jones, C. K. In Multiphase polymer materials: blends, ionomers and interpenetrating networks; Utracki, L. A., Weiss, R. A., Eds.; ACS Symposium Series; American Chemical Society: Washington, DC, 1989; pp 401-417. (39) Sakai, T.; Takenaka, H.; Torikai, E. J. Membr. Sci. 1987, 31, 227-234.

Figure 6. Conductivity of the effluents from the capture device during a run. In A-F, the run conditions and the time that the capture voltage was turned off (arrows) were the same as those described in Figure 4A-F. In G, the polarity of the electric field was reversed at 1200 s (inverted arrow). Curves were displaced along the y axis for clarity.

far less. At its peak, the recovered DNA band was ∼12 times more concentrated than the sample. Greater concentration factors can likely be obtained by increasing the volume of sample loaded or by shortening the length of outlet tubing. We have generally observed that increasing the volume of sample applied did not significantly reduce the recovery of DNA. Desalting and Sample Cleanup. In capillary electrophoresis, salt and buffer ions (for example, from a DNA sequencing or other enzymatic reaction) will drastically reduce the amount of sample loaded and, hence, sensitivity and can also cause deterioration in separation quality.16-18,40-42 Therefore, to be useful as a general method for sample cleanup, desalting the DNA during capture is highly desirable. After capturing DNA fragments, washing the captured DNA with deionized water for 10 min typically resulted in the removal of >90% of salt and buffer ions from the recovered DNA samples. Figure 6 shows the conductivity of the material recovered from the capture device, monitored at its outlet. High conductivity was evident prior to capture of the DNA fragments from the sample solution (signal near 0 min). The salt concentration in samples A-C was drastically reduced during extended washes with deionized water (Figure 6D-G). After an initial fast reduction, salt concentration continued to decrease for some time, but beyond this, a further reduction in the salt concentration was not observable with prolonged washing. The area under the conductivity curve for the eluted band after a 10-min wash was ∼4-8% of that for an uncaptured sample band (compare curves A and F in Figure 6). Application of a reversedpolarity electric field did not increase salt elution from the device (curve G, Figure 6). A substantial difference in electrophoretic (40) Schwartz, H. E.; Ulfelder, K.; Sunzeri, F. J.; Busch, M. P.; Brownlee, R. G. J. Chromatogr. 1991, 559, 267-283. (41) Tong, X.; Smith, L. M. Anal. Chem. 1992, 64, 2672-2677. (42) Figeys, D.; Ahmadzedeh, H.; Arriaga, E.; Dovichi, N. J. J. Chromatogr., A 1996, 744, 325-331.

Figure 7. Signal amplification associated with application of the capture device. A portion of a conventional capillary electrophoresisbased DNA sequencing run is shown. CE was performed on an ABI 310 system under identical conditions for the four sample preparations, all TAMRA-labeled M13mp18 “G” sequencing reactions. A, crude reaction products; B, purified reaction products (purification using conventional ethanol-precipitation); C, 3 µL of crude reaction products concentrated with the capture device; D, 10 µL of crude reaction products concentrated with the capture device. Capture conditions are described in the Experimental Section.

mobility and membrane permeability of DNA and salt is likely responsible for the difference in recovery ratios (95% vs