Anal. Chem. 2003, 75, 1218-1220
Technical Notes
Effect of Caged Fluorescent Dye on the Electroosmotic Mobility in Microchannels David Ross*,† and Laurie E. Locascio‡
Process Measurement Division and Analytical Chemistry Division, National Institute of Standards & Technology, Gaithersburg, Maryland 20899
We report on measurements of electroosmotic mobility in polymer microchannels and silica capillaries with and without the addition of a caged fluorescein dye to the buffer. For PMMA microchannels, the mobility was found to increase from (2.6 ( 0.1) × 10-4 cm2 V-1 s-1 to (4.6 ( 0.1) × 10-4 cm2 V-1 s-1 upon addition of 1.2 mmol/L of caged dye. For PC microchannels, the mobility increased from (4.3 ( 0.2) × 10-4 cm2 V-1 s-1 to (5.4 ( 0.1) × 10-4 cm2 V-1 s-1 upon addition of caged dye. For PDMS microchannels, the mobility increased from (4.3 ( 0.2) × 10-4 cm2 V-1 s-1 to (6.4 ( 0.5) × 10-4 cm2 V-1 s-1 upon addition of caged dye. For fused-silica capillaries, the mobility ((5.5 ( 0.2) × 10-4 cm2 V-1 s-1) was unaffected by the addition of the caged dye. A number of recent papers in Analytical Chemistry have reported on the use of caged fluorescent dyes1 for the measurement of electroosmotic flow (EOF) in microfluidic systems.2-6 Caged dyes are fluorescent dyes that have been made nonfluorescent through the binding of chemical groups. These “caging” groups can be removed by exposing the caged dye to UV light, restoring the dye to its uncaged (and fluorescent) state. In a typical caged dye experiment, a focused and spatially localized UV laser pulse is used to uncage the dye in a selected portion of the device being tested, thereby marking the flow. The use of caged dyes has a number of advantages for flow measurement in microfluidics, including the high signal-to-noise ratio inherent to fluorescencebased techniques and the lack of perturbation of the flow as the marker is created in situ. However, the technique is not without its drawbacks. As yet, the available, water-soluble caged dyes are †
Process Measurement Division. Analytical Chemistry Division. (1) Lempert, W. R.; Magee, K.; Ronney, P.; Gee, K. R.; Haugland, R. P. Exp. Fluids 1995, 18, 249-257. (2) Paul, P. H.; Garguilo, M. G.; Rakestraw, D. J. Anal. Chem. 1998, 70, 24592467. (3) Herr, A. E.; Molho, J. I.; Santiago, J. G.; Mungal, M. G.; Kenny, T. W.; Garguilo, M. G. Anal. Chem. 2000, 72, 1053-1057. (4) Molho, J. I.; Herr, A. E.; Mosier, B. P.; Santiago, J. G.; Kenny, T. W.; Brennen, R. A.; Gordon, G. B.; Mohammadi, B. Anal. Chem. 2001, 73, 1350-1360. (5) Ross, D.; Johnson, T. J.; Locascio, L. E. Anal. Chem. 2001, 73, 25092515. (6) Johnson, T. J.; Ross, D.; Gaitan, M.; Locascio, L. E. Anal. Chem. 2001, 73, 3656-3661. ‡
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all charged when uncaged so that the results must be corrected for the electrophoretic motion of the dye after uncaging. To further complicate this correction, multiple fluorescent species with different electrophoretic mobilities can result from uncaging of a single dye.7 In addition, as we report here, the use of caged dye in a microfluidic system can alter the EOF mobility through nonspecific adsorption of the caged dye to the walls of the microsystem. This effect is particularly pronounced in polymer substrate systems and can result in large errors in EOF measurements. EXPERIMENTAL SECTION Chemicals and Materials. Fluorescein bis(5-carboxymethoxy2-nitrobenzyl) ether dipotassium salt (CMNB-caged fluorescein dye) was used as supplied by Molecular Probes8 (Eugene, OR). Buffer solutions were made using deionized water from a Millipore Milli-Q8 system (Bedford, MA). A solution of 200 mM carbonate buffer, pH 9.4, was prepared according to instructions using a carbonate-bicarbonate buffer pack (Pierce,8 Rockford, IL) and was diluted to make the 18- and 20-mM buffers used for the current monitoring measurements. Solutions of 1.2 mM caged dye were prepared by dissolving 5 mg of CMNB-caged fluorescein solid in 5 mL of the carbonate buffer and were filtered before use with syringe filters. Microchannels were made using 3-mm-thick poly(methyl methacrylate) (PMMA) sheet (Acrylite OP-4,8 35 Cyro Industries,8 Mt. Arlington, NJ), 125-µm-thick polycarbonate (PC) sheet (McMaster-Carr, Atlanta, GA), or poly(dimethylsiloxane) (PDMS). Fused-silica capillary tubing (360-µm o.d., 50-µm i.d.) was obtained from Polymicro Technologies Inc.8 (Phoenix, AR). Poly(dimethylsiloxane) (PDMS) was prepared according to product information from a Sylgard 1848 silicone elastomer kit (Dow Corning,8 Midland, MI) and cured at room temperature for 1 week. Microchannel Fabrication. Polymeric microchannels were formed by imprinting with a micromachined silicon template and then sealing with similar polymeric material as has been previously (7) Molho, J. I. Ph.D. Thesis, Stanford University, Stanford, CA, 2001. (8) Certain commercial equipment, instruments, or materials are identified in this report to specify adequately the experimental procedure. Such identification does not imply recommendation or endorsement by the National Institute of Standards and Technology, nor does it imply that the materials or equipment identified are necessarily the best available for the purpose. 10.1021/ac026277u CCC: $25.00
© 2003 American Chemical Society Published on Web 01/24/2003
described.5,9 Typical microchannels were trapezoidal in cross section and were 75 µm wide at the base, 25 µm wide at the top, and 30 µm deep. PMMA and PC microchannels were imprinted into rectangular pieces (23 × 38 mm) of the polymer sheet material. PMMA channels were imprinted for 20 min at 110 °C under an applied force of 4500 N and sealed to a sheet of similar PMMA at 103 °C for 10 min in a circulating air oven. PC channels were imprinted at 155 °C at 13 000 N and then cooled to 120 °C and released. The PC channels were then sealed to a similar sheet of PC at 135 °C for 1 h. PDMS channels were formed by pouring a Sylgard 1848 silicone elastomer mixture onto the silicon template and allowing it to cure at room temperature for ∼1 week. The PDMS channels were then sealed with a flat sheet of cured PDMS from the same batch as was used for the channels. PDMS channels were not modified in any way after curing. Fused-silica capillaries were stripped of their polyimide coating and embedded in the middle of a sheet of PDMS to provide an object geometrically similar to the plastic microchannels. The channels and capillaries used were each ∼2 cm long. Electroosmotic Mobility Measurements. The electroosmotic mobility was determined using an automated version of the current monitoring method10 that is described in detail elsewhere.11 Briefly, the microchannel and one of the fluid reservoirs were filled with buffer of one concentration (18 mM), and the second fluid reservoir was filled with a similar buffer of higher concentration (20 mM). High voltage was applied, and the currentmonitoring method was used to measure the EOF velocity as the 20 mM buffer filled the channel, displacing the 18 mM buffer. After the 20 mM buffer had completely filled the channel, the voltage was turned off for 60 s to allow the buffer in the first reservoir to relax to a uniform concentration, after which the voltage was reapplied with the polarity reversed so that EOF was driven from the first reservoir toward the second. The current was again monitored as a function of time as the 18 mM buffer moved into the channel, displacing the 20 mM buffer. This procedure of switching the polarity of the applied voltage was repeated several times at two different applied field strengths (100 V/cm and 200 V/cm), with the voltage turned off for 60 s each time the polarity was reversed. The current measurements and the switching of the voltage were automated and computercontrolled. RESULTS AND DISCUSSION For each microchannel used, the electroosmotic mobility was determined three times using the current-monitoring method as described above. First, the mobility of the channel or capillary was measured using carbonate buffer without caged dye. Then, the channel was filled with buffer containing 1.2 mM caged dye, and the measurements were repeated to determine the effect of nonspecific adsorption of the caged dye. Finally, the fluid reservoirs were rinsed with buffer (no caged dye), 3-5 µL of buffer was flushed through the channel, and the measurements were repeated again without dye to see if the effects of nonspecific adsorption were reversible. (9) Martynova, L.; Locascio, L. E.; Gaitan, M.; Kramer, G. W.; Christensen, R. G.; MacCrehan, W. A. Anal. Chem. 1997, 69, 4783-4789. (10) Huang, X. H.; Gordon, M. J.; Zare, R. N. Anal. Chem. 1988, 60, 18371838. (11) Ross, D.; Henry, A. C. to be published.
Table 1. Measured EOF Mobilities for PMMA, PC, PDMS, and Silica with 18/20 mM Carbonate Buffer pH 9.4
PMMA PDMS PC silica
without caged dye 10-4 cm2/V/sa
with caged dye 10-4 cm2/V/sa
rinsed 10-4cm2/V/sa
2.6 ( 0.1 4.3 ( 0.1 4.3 ( 0.2 5.6 ( 0.3
4.6 ( 0.1 6.4 ( 0.5 5.4 ( 0.1 5.4 ( 0.1
3.9 ( 0.1 6.5 ( 0.1 5.2 ( 0.1 5.5 ( 0.1
a Column 1, clean channels with no caged dye added to buffer. Column 2, the same channels but with 1.2 mM caged dye added to buffer. Column 3, the same channels again, after rinsing with buffer, and measurement made with no caged dye added to buffer. In each case, the values and uncertainties given are the average and standard deviation of a set of at least seven repeated measurements on a single channel.
Figure 1.
The results are summarized in Table 1. The uncertainties given are the one-standard-deviation statistical uncertainties resulting from each set of repeated measurements on a single microchannel. Device-to-device variability can be significantly greater than the given uncertainties, particularly if plastics from different vendors are used. Additional systematic uncertainties are estimated to be comparable to or less than the listed statistical uncertainties. The values obtained without caged dye are comparable to those reported in the literature for similar conditions for PMMA,12 PDMS,13 PC,14 and silica.15 For the polymeric microchannels, the addition of caged dye significantly increased the electroosmotic mobility, with particularly large increases for PMMA and PDMS. For PMMA, the mobility increase is partiallysbut not completelysreversible after rinsing the channels with buffer. The electroosmotic mobility in fused-silica capillaries was not significantly affected by the addition of caged dye. Considering the structure of the caged-dye molecule (Figure 1), it is not surprising that a significant degree of adsorption could be found for plastic surfaces and not for silica. The adsorption is likely due to nonspecific hydrophobic interactions between the plastic substrate and the aromatic moieties of the caged dye molecule. Further, because of the two anionic charge groups of the caged dye molecule, adsorption would be expected to result in an increase in the EOF, as was observed experimentally. This kind of behaviorsa large change in EOF resulting from relatively (12) Henry, A. C.; Tutt, T. J.; Galloway, M.; Davidson, Y. Y.; McWhorter, C. S.; Soper, S. A.; McCarley, R. L. Anal. Chem. 2000, 72, 5331-5337. (13) Ocvirk, G.; Munroe, M.; Tang, T.; Oleschuk, R.; Westra, K.; Harrison, D. J. Electrophoresis 2000, 21, 107-115. (14) Soper, S. A.; Henry, A. C.; Vaidya, B.; Galloway, M.; Wabuyele, M.; McCarley, R. L. Anal. Chim. Acta 2002, 470, 87-99. (15) Harrison, D. J.; Manz, A.; Fan, Z. H.; Ludi, H.; Widmer, H. M. Anal. Chem. 1992, 64, 1926-1932.
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low concentration of an additive to the buffersis not unprecedented for polymeric microchannels and has been found for the addition of both anionic13,16 and cationic17 surfactants to the buffer. CONCLUSIONS The results presented here indicate a potential problem with the caged-dye method of flow imaging. Altough the method is relatively nonperturbative in that the flow marker is created in situ without interruption of the flow, the adsorption of the caged dye onto the walls of the system can dramatically affect the results, particularly for electrokinetically driven flow. Fortunately, the effects of caged dye on the electroosmotic mobility seem to be confined to polymer substrate systems, though this should be verified with each particular substrate material. The results presented here bring into doubt the quantitative accuracy of previously reported measurements of EOF in plastic microchannels obtained using caged dye imaging.5 The EOF mobility results of the previous report for plastic microchannels (16) Xu, W.; Uchiyama, K.; Shimosaka, T.; Hobo, T. J. Chromatogr., A 2001, 907, 279-289. (17) Wang, S. C.; Perso, C. E.; Morris, M. D. Anal. Chem. 2000, 72, 17041706.
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are consistent with the results obtained here with caged dye added, indicating that nonspecific adsorption of the caged dye was probably occurring then as well. Consequently, we can infer that EOF measurements obtained using the caged dye technique would be in error by approximately 75% for PMMA and 50% for PDMS. Nevertheless, the qualitative conclusions of the previous paper concerning the use of combinations of different materials in the construction of microchannels and the effects of nonuniform ζ potential on sample dispersion under EOF (see also refs 3,6) are likely to be correct, although the magnitude of the variations in ζ potential might have been influenced by the presence of the caged dye. This result clearly highlights the need for the development of improved probes and methods for detailed flow measurement in more complex microfluidic systems with nonuniform ζ potential and made with a variety of materials, particularly polymers.
Received for review December 19, 2002. AC026277U
October
31,
2002.
Accepted
