- Email: [email protected]
Thermoplastic microchannel fabrication using carbon dioxide laser ablation
Journal of Chromatography A, 1111 (2006) 252–257
Thermoplastic microchannel fabrication using carbon dioxide laser ablation Shau-Chun Wang ∗ , Chia-Yu Lee, Hsiao-Ping Chen 160 San-Hsing, Ming-Hsiung, Department of Chemistry and Biochemistry, National Chung Cheng University, Chia-Yi 621, Taiwan Available online 8 November 2005
Abstract We report the procedures of machining microchannels on Vivak co-polyester thermoplastic substrates using a simple industrial CO2 laser marker. To avoid overheating the substrates, we develop low-power marking techniques in nearly anaerobic environment. These procedures are able to machine microchannels at various aspect ratios. Either straight or serpent channel can be easily marked. Like the wire-embossed channel walls, the ablated channel surfaces become charged after alkaline hydrolysis treatment. Stable electroosmotic flow in the charged conduit is observed to be of the same order of magnitude as that in fused silica capillary. Typical dynamic coating protocols to alter the conduit surface properties are transferable to the ablated channels. The effects of buffer acidity on electroosmotic mobility in both bare and coated channels are similar to those in fused silica capillaries. Using video microscopy we also demonstrate that this device is useful in distinguishing the electrophoretic mobility of bare and latex particles from that of functionalized ones. © 2005 Elsevier B.V. All rights reserved. Keywords: Microfabrication; Microfluidics; Electrophoretic mobility; Plastics
1. Introduction Polymeric materials are attractive substrates for fabrication of microfluidics devices. Fabrication techniques appropriate to several important polymeric materials have been described. These polymers include thermoplastics such as poly(methyl methacrylate) (PMMA), polycarbonate and co-polyesters, and elastomers such as poly(dimethylsiloxane) (PDMS) [1]. In addition, polymer-based devices have also been used to distinguish cells or particles based on their electrophoretic mobility difference [2,3]. The fabrication technologies to machine silicon and glass chips, including photolithography, plasma etching, laser/X-ray ablation [4], have also been employed on plastic chips. Simple methods such as wire and template imprinting and injection molding are also suitable for fabricating polymeric materials, especially engineering plastics [1]. This technological progress suggests that silica or glass chip can be replaced with polymeric materials, with apparent advantages that are associated with simple fabrication on soft channels. But the initial development of such molds still requires considerable effort and expense.
∗
Corresponding author. Tel.: +886 5 2720411x66410; fax: +886 5 2721040. E-mail address: [email protected] (S.-C. Wang).
0021-9673/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2005.10.039
Recently, machining microchannel devices with simple industrial CO2 laser marker on plastic substrates such as PMMA have been reported for the applications of nucleic acid hybridization and optical sense using [5,6]. These laser markers have reasonable cost (20k US dollars or less) and are capable of easily fabricating typical two-dimensional micro-structures. Besides, no extra cost of either hardware or software development is necessary to fabricate microchannels at moderately accelerated throughput of approximately tens of chips per day. The production scale and initial cost fits the demand of a bio-analytical laboratory, wherein versatility is equally important as economical advantages in developing tens of chips for preliminary evaluation. To sustain adequate electroosmotic flows in microfluidic devices for fluid pumping, surface treatment on PMMA channel walls using strong reduction reagents has been studied [5]. The use of UV lasers to initiate photochemical reactions on plastics surface is an alternative protocol to generate electroosmotic flows in PMMA microchannel by charging the initially neutral acrylic substrate [7]. In contrast, generating electroosmotic flows in co-polyester microchannels is much easier. Ester functionality produces charged carboxylate group via hydrolysis when the channel is filled with diluted basic or acidic buffers [8,9]. The acidity dependence of electroosmotic mobility in co-polyester channels confirms the above mechanism [10]. Although
S.-C. Wang et al. / J. Chromatogr. A 1111 (2006) 252–257
co-polyester substrate absorbs UV and blue light considerably, this optical interference problem is curable with confocal detection, polarized fluorescence detection, or analyte velocity modulation method [11–14]. In this paper, we use a simple industrial CO2 laser marker to craft microchannels on Vivak co-polyester chips. Built-in pattern-marking software is used to control the writing motions of a low-power laser beam. To avoid overheating the plastic substrate, dry ice is placed in the marking chamber to significantly suppress the partial pressure of oxygen. Because the laser power is low, the laser beam has to move repetitively along the same line on a chip for several times to write one microchannel. The typical channel width is 300–350 m. The number of repetitive writing controls the aspect ratio of laser-ablated microchannel. We have measured the electroosmotic mobility in a copolyester microchannel after hydrolyzing it with buffers. The mobility is almost the same as that in the wire-imprinted microchannels reported previously [9,10]. The mobility is also stable whose measurement variation is less than 0.2%. Using surfactant cetyltrimethylammonium bromide (CTAB), we also confirm that dynamic coating protocols in silica capillaries are transferable to laser-marked plastic microchannels. Using video microscopy to measure the electrophoretic mobility of latex particles in the microchannel, we obtain the mobility histograms of particle mixture showing clear difference between bare and fluoroscein-tagged particles. 2. Experimental 2.1. Materials Co-polyester plastic slides are cut from Vivak plastic sheets (DSM, Sheffield, MA, USA). ACS grade reagents, including boric acid, sodium carbonate, acetic acid, citric acid, and sodium hydroxide, are used for the preparation of buffers. Cetyltrimethylammonium bromide is obtained from Sigma (St. Louis, MO, USA). We purchase bare (5 m) and fluorescein-tagged (10 m) latex particles from Fluka (Switzerland) and Polysciences (PA, USA), respectively.
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the repetition count controls the aspect ratio of laser-ablated microchannel. To inspect the marked channel from the top view, optical microscopy is employed. The cross section of a channel is imaged using scanning emission microscopy (S-2400, Hitachi, Japan). 2.3. Measurement of electroosmotic mobility in microchannel We flush Tris–Borate–EDTA (TBE) buffer through our lasermarked microchannel to hydrolyze co-polyester substrate on the wall into the charged carboxylate functionalities. These charged moieties are able to establish an electrical double layer within the electrolyte solution. Electroosmotic flows are then produced when an electric field (50–400 V/cm) is applied across the channel. A current monitoring method is adopted to measure the current transition duration time during the replacement of a higher conductivity buffer with a lower one using electroosmosis [15]. Typically, dilute buffer is filled in the cathode reservoir to aspirate into the uncoated channel using electroosmosis after the electrophoresis voltage is turned on, and the dilute buffer gradually takes over the higher concentration buffer filled in the channel. Current through the channel is recorded with the voltage drop across a high resistor between the anode electrode and the ground. We employ four types of buffer solutions, sodium citrate, sodium acetate, sodium carbonate, and sodium borate, all at 25 mM to measure electroosmotic mobility at pH 3, 5, 7, and 9, respectively. Since the electroosmotic mobility is linearly proportional to the charge density on the channel surface (Helmholtz–Smoluchowski equation), the relative ratios of negative charge density on the channel surface between these pH values are obtained from the mobility data. The mobility values in CTAB-coated channels are also obtained using the current monitoring method at the same pH conditions as those described above. Since the surface of coated channel is positively charged, the direction of the electroosmotic flow is opposite to that in uncoated channel. Therefore, dilute buffer is filled in the anode reservoir to aspirate into the channel to replace higher concentration buffers.
2.2. Fabrication of microchannels We use an industrial CO2 laser marker (Model TL-25P, TopDog Laser, Taiwan) to fabricate microchannels on co-polyester chips. The laser beam has a wavelength of 10.6 m and the focus spot is approximately 100 m in diameter. The lasing tube is housed in a rack, whose motion is controlled by the buildin software. The housing rack is free to move on a horizontal surface in a chamber (70 cm × 60 cm × 26 cm) to strike the laser beam within an area of 24 cm × 24 cm. The writing speed of the laser beam is 15 cm/s. Approximately 100 g of dry ice is placed in the marking chamber to significantly dilute the partial pressure of oxygen. During the marking procedures, the laser power is intentionally reduced to 1.4 W to avoid overheating the chip. We control the laser beam to move repetitively along the same line on the chip for several times to machine one microchannel;
2.4. Observation of latex particle migration in microchannel to distinguish electrophoretic mobility We use a microscope to trace the movement of bare and fluorescein-tagged latex particles in the microchannel. Treating the microchannel surface with borate (25 mM) buffer, we apply an electric field of 75 V/cm across the channel filled with borate (25 mM). The concentrations of bare and functionalized latex particles seeded in the microchannel are 9.1 × 105 and 1.82 × 105 particles/ml, respectively. Although the particles are neutral or negatively charged, they move toward the ground electrode due to the electroosmotic flow. With a CCD camera (DFW-V500, Sony, Japan) operating at the acquisition rate of 15 frame/s, we acquired the images of particle movement under the focus region of an objective lens on the Nikon microscope.
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Particle trajectory and speed is analyzed with image processing software (ACDSee 6.0, ACD Systems, Canada). 3. Results and discussion Fig. 1A and B show the photographic images of the CO2 laser-marked channels on co-polyester slides using the proposed fabrication procedures. Fig. 1A shows a straight channel fabricated with the protocols described in Section 2. The serpent channels illustrated in Fig. 1B, on the other hand, can be fabricated by first machining the horizontal channels and later marking the connecting vertical channels with similar protocols. The widths of these channels are between 300 and 350 m. Therefore, the proposed procedures are demonstrated to be capable of marking straight or serpent channel. Fig. 2 shows the SEM picture of a cross-section of a laser-marked channel with onehalf aspect ratio. The channel surface can be observed to be adequately smooth. The SEM pictures (not shown) of channels at different aspect ratio (approximately 0.5–2.0) indicate similar smoothness.
Fig. 2. SEM image depicting the cross-section of a co-polyester microchannel created with CO2 laser marker. The channel aspect ratio is 0.53.
We used current monitoring method to measure electroosmotic mobility in the single straight channel chip (Fig. 1A). A typical curve showing the process of replacing the concentrated buffer (sodium acetate; 25 mM) with a dilute one (20 mM) is illustrated in Fig. 3. The current across the channel gradually drops while the dilute buffer is being filled into the channel. Except at the beginning and the end of the buffer replacement process, this current drop curve is almost linear. The regression result using the data of linear curve shows the correlation coefficient (r) is 0.997. The standard deviation of residue values (i.e. standard error) is 3.08 × 10−4 mA, which is less than 0.2% of the average current level (2.66 × 10−1 mA) of 25 and 20 mM buffer. Since the electroosmotic mobility variation during the buffer displacement process in the channel is virtually negligible, the mobility is therefore proved stable. Although the channels are not perfectly straight, slight irregularities in the channel do not influence the electroosmotic flow stability, as will be subsequently discussed. We also measure electroosmotic mobility in three different single channel chips (Fig. 1A) cut from the same
Fig. 1. (A) Single segment of a straight microchannel fabricated on a copolyester chip with CO2 laser marker. (B) Co-polyester substrate fabricated with a serpent channel using CO2 laser marker. The channel width in parts A and B is 324 m. The channel bottoms have been dyed with ink to improve visualization.
Fig. 3. The channel current monitored in the process of displacing 25 mM acetate buffer solution with 20 mM solution. The straight line shows the linear regressed curve using the data in the middle of displacement process.
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Fig. 4. The effect of buffer pH values on electroosmotic mobility in hydrolyzed and surfactant-coated co-polyester microchannels. The upper trace shows the buffer effect in a bare channel. The electroosmotic mobility in pH 3 is too small to detect. The lower trace shows the effect in a coated channel. The coating reagent is cetyltrimethylammonium bromide (CTAB).
plastic sheet. These mobility values were found to be 2.9 (±0.15) × 10−4 cm2 /V s in acetate buffer (pH 4.8). This negligible variation (less than 3.6%) in the measured mobility is consistent with the reported value in wire-imprinted channel [9,10]. We illustrate the pH dependence on electroosmotic mobility data measured with the current monitoring method in bare hydrolyzed co-polyester microchannels and surfactant-coated ones in Fig. 4. The curve for the bare channel keeps increasing as buffer acidity changes from acidic to neutral and becomes almost constant above pH 6. The other curve for the CTABcoated channel indicates the similar pH dependence of electroosmotic mobility to that in hydrolyzed co-polyester microchannels. Although the direction of electroosmotic flows in coated channels is reversed, the flow rate also increases from acidic to neutral pH conditions and become constant. The trends of these two curves are similar to those in previous reports using wire-imprinted channels [9,10]. Fig. 5 shows five consecutive images tracing the migration of bare latex particles seeded in the microchannel filled with sodium borate buffer (25 mM; pH 9). The positions of three traced particles driven in the electric field 75 V/cm are highlighted by circle, square, and oval marks. The outward arrows from the marks show the direction of the particle as it translates. Clearly, the migration of these particles is unidirectional suggesting the absence of any circulation in the bulk flow due to variations in the electroosmotic flow on the channel wall [16]. This observation indicates the negligible mobility variation in the channel, which is consistent with the conclusion obtained with current monitoring method in the above. Theoretically, the stability of electroosmotic flow rates is not sensitive to the variation of channel width when the charge density of the channel wall is homogenous [17]. Since the hydrolysis conversion of co-polyester to carboxylate takes place in the channel without spatial preference, the wall charge density distribution has to be uniform. When the characteristic length of channel is much larger than the Debye length, typically 10–100 nm in dilute electrolyte solutions, the Smoluchowski slip
Fig. 5. Five consecutive image snapshots taken once a second showing the migration trajectories of three latex particles moving across one 1500 m zone under the microscope objective. The traced particles are indicated with circle, square, and oval marks, respectively. The horizontal arrows show the migration direction of particles. The channel is filled with sodium borate buffer (25 mM; pH 9). The electric field applied across the channel is 75 V/cm.
velocity, which is proportional to the tangential field, applies at the slip plane. Since there is normal velocity on the surface, a continuous surface is then covered by the streamlines of the flow. As the surface charge screens the external field, the electric field is exactly tangential to the charged surface anywhere. Moreover, the governing equations for the interstitial flow driven by the slip velocity on the surface is the Laplace equation for the velocity potential, which is identical to the Laplace equation for electric potential. Such duality of boundary conditions and governing equations for the velocity potential and electric potential implies that the electric field lines and the streamlines are identical in the bulk if the surface charge is uniform. This has been proven by Takhistov et al. [18]. As a consequence of this duality, the flow rate is identical to the current for a channel with uniform charge. Another corollary is that, while the total flow rate is dependent on the transverse geometry and dimension of the channel, the electroosmotic mobility and local velocity are not. Our electroosmotic mobility measurement data in ablated co-polyester channel show almost equivalent mobility to that in fused-silica capillary. This result also suggests that the channel surface is smooth, not coarse. With uneven roughness, the local slip velocity would not be uniform and pressure-driven back flow would result producing vortices and transverse particle trajectories such as those observed by Minerick et al. [16] and Takhistov et al. [18]. Our data show that the unidirectional longitudinal particle migration driven by electroosmosis and the steadiness of buffer displacement process in the channel are both consistent with our theoretical conjecture that the channel is smooth.
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Although electro-osmotic mobility in the CO2 laser-ablated channel is steady, capillary electrophoresis (CE) separations in the channel is not feasible because the channel width (300–350 m) is not narrow enough. The plastic conduit fabricated with the laser marker is somewhat short of surface force to resist the hydrostatic pressure across the channel. The surface force is proportional to the channel width but the hydrostatic force is proportional to the channel cross-section (the square of channel width). When channel width increases, the hydrostatic force exceeds the surface force. Therefore, analyte loading
becomes questionable for sustaining limited dispersion of sample zone. Since the channel mobility is stable, particles with different electrophoretic mobility are distinguishable in the channel. Using particle trajectory data acquired with video microscopy, particle mobility values can be obtained. Fig. 6A and B show the individual mobility histograms of bare and fluorescein-tagged latex particles in the microchannel, respectively. The inset figures are the simulated electropherograms, whose peaks are depicted using the mean values and standard deviations obtained as shown in Fig. 6A and B. When the channel is filled with borate buffer (25 mM; pH 9.2), its electroosmotic mobility of 6.4 × 10−4 cm2 /V s is determined. The surface charge of bare latex particle is usually small. Therefore, the apparent mobility of bare particles in the hydrolyzed channel should be very close to the electroosmotic mobility of channel. The apparent mobility of bare particle on average is estimated as 6.4 × 10−4 cm2 /V s. This mobility value is reasonable. On the other hand, fluoresceintagged particles are more negatively charged. The mean apparent mobility of these functionalized particles should be somewhat less than the channel mobility, which is consistent with the result, 3.7 × 10-4 cm2 /V s, determined from the histogram of Fig. 6B. The Fig. 6C shows the mobility histogram of particle mixture in the same channel. This histogram has two distinct clusters, whose mean mobility values, 6.4 × 10−3 and 3.7 × 10−4 cm2 /V s, respectively, are identical to those estimated in Fig. 6A and B. The peaks in the simulated electrophoregrams in the inset figures are depicted using the mean values and standard deviations of two clusters in Fig. 6C. Having been easily classified as two distinct clusters, the mobility values of two bare and functionalized latex particles are distinguishable in the laser-ablated co-polyester channel. 4. Conclusions
Fig. 6. The electrophoretic mobility histograms in the channel of bare latex particles frame A, fluorescein-tagged latex particles frame B, and the mixtures of two types of latex particles frame C. The channel is filled with borate buffer solution (25 mM). The insets in A–C show the simulated electropherograms converted from the mobility histograms.
We have successfully developed a technique to fabricate microchannels on co-polyester substrates using an industrial CO2 laser marker. When the laser power is weak and partial pressure of oxygen in the machining chamber has been suppressed, repetitive writing motions of the laser beam on the chip gradually machine a microchannel of 300–350 m width. By controlling the writing power and repetition times, we are able to fabricate the microchannel at varied aspect ratios from approximately 0.5–2. Similar to wire-imprinted channels, laser-ablated copolyester channels undergo hydrolysis to become charged. Although the channel width is not perfectly consistent, stable electroosmotic mobility is validated in the laser-ablated channel. In addition, we show that dynamic coating protocols in capillary electrophoresis are transferable to the laser-marked channel. This channel width is too wide to provide adequate surface force to avoid solute dispersion in the sample loading process for performing capillary electrophoresis separations. By using video microscopy we demonstrate that CO2 laser-ablated channel is suitable to distinguish bare latex particles from functionalized ones with the particle mobility histograms.
S.-C. Wang et al. / J. Chromatogr. A 1111 (2006) 252–257
Acknowledgements The authors express thanks for the financial support from National Science Council, Taiwan (NSC92-2113-M-194-022 and NSC93-2120-M-194-005), and National Chung Cheng University. The authors also acknowledge helpful discussions with L.Y. Yeo and H.-C. Chang (Department of Chemical and Biomolecular Engineering, University of Notre Dame, IN, USA) in the preparation of this manuscript. We also thank J.-Y. Cheng (Center for Applied Sciences, Academia Sinica, Taiwan) for discussions on laser-marking techniques. References [1] T.D. Boone, Z.H. Fan, H.H. Hooper, A.J. Ricco, H. Tan, S.J. Williams, Anal. Chem. 74 (2002) 78A. [2] A.Y. Fu, C. Spence, A. Scherer, F.H. Arnold, S.R. Quake, Nat. Biotechnol. 17 (1999) 1109. [3] M.A. Witek, S.Y. Wei, B. Vaidya, A.A. Adams, L. Zhu, W. Stryjewski, R.L. McCarley, S.A. Soper, Lab on a Chip 4 (2004) 464.
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[4] M. Madou, Fundamentals of Microfabrication, CRC Press, Boca Raton, FL, USA, 1997. [5] J.-Y. Cheng, C.-W. Wei, K.-H. Hsu, T.-H. Young, Sens. Actuators B 99 (2004) 186. [6] H. Klank, J.P. Kutter, O. Geschke, Lab on a Chip 2 (2002) 242. [7] T.J. Johnson, D. Ross, M. Gaitan, L.E. Locascio, Anal. Chem. 73 (2001) 3656. [8] L.E. Locascio, C.E. Perso, S.L. Cheng, J. Chromatogr. A 857 (1999) 275. [9] S.-C. Wang, M.D. Morris, Anal. Chem. 72 (2000) 1704. [10] S.-C. Wang, C.-Y. Lee, H.-P. Chen, Biosens. Bioelectron. 20 (2005) 2126. [11] S.-C. Wang, M.D. Morris, J. Chin. Chem. Soc. 52 (2004) 759. [12] S.-C. Wang, C.E. Perso, M.D. Morris, Anal. Chem. 72 (2000) 1448. [13] S.-C. Wang, Biosens. Bioelectron. 20 (2004) 139. [14] S.-C. Wang, H.-P. Chen, C.-Y. Lee, Biosens. Bioelectron. 20 (2005) 2131. [15] X. Huang, M.J. Gordon, R.N. Zare, Anal. Chem. 60 (1988) 1837. [16] A. Minerick, A. Ostafin, H.-C. Chang, Electrophoresis 23 (2002) 2165. [17] K.P. Tikhomolova, Electroosmosis, Harwood, New York, 1993. [18] P. Takhistov, K. Duginova, H.-C. Chang, J. Colloid Interface Sci. 263 (2003) 133.
Thermoplastic microchannel fabrication using carbon dioxide laser ablation Shau-Chun Wang ∗ , Chia-Yu Lee, Hsiao-Ping Chen 160 San-Hsing, Ming-Hsiung, Department of Chemistry and Biochemistry, National Chung Cheng University, Chia-Yi 621, Taiwan Available online 8 November 2005
Abstract We report the procedures of machining microchannels on Vivak co-polyester thermoplastic substrates using a simple industrial CO2 laser marker. To avoid overheating the substrates, we develop low-power marking techniques in nearly anaerobic environment. These procedures are able to machine microchannels at various aspect ratios. Either straight or serpent channel can be easily marked. Like the wire-embossed channel walls, the ablated channel surfaces become charged after alkaline hydrolysis treatment. Stable electroosmotic flow in the charged conduit is observed to be of the same order of magnitude as that in fused silica capillary. Typical dynamic coating protocols to alter the conduit surface properties are transferable to the ablated channels. The effects of buffer acidity on electroosmotic mobility in both bare and coated channels are similar to those in fused silica capillaries. Using video microscopy we also demonstrate that this device is useful in distinguishing the electrophoretic mobility of bare and latex particles from that of functionalized ones. © 2005 Elsevier B.V. All rights reserved. Keywords: Microfabrication; Microfluidics; Electrophoretic mobility; Plastics
1. Introduction Polymeric materials are attractive substrates for fabrication of microfluidics devices. Fabrication techniques appropriate to several important polymeric materials have been described. These polymers include thermoplastics such as poly(methyl methacrylate) (PMMA), polycarbonate and co-polyesters, and elastomers such as poly(dimethylsiloxane) (PDMS) [1]. In addition, polymer-based devices have also been used to distinguish cells or particles based on their electrophoretic mobility difference [2,3]. The fabrication technologies to machine silicon and glass chips, including photolithography, plasma etching, laser/X-ray ablation [4], have also been employed on plastic chips. Simple methods such as wire and template imprinting and injection molding are also suitable for fabricating polymeric materials, especially engineering plastics [1]. This technological progress suggests that silica or glass chip can be replaced with polymeric materials, with apparent advantages that are associated with simple fabrication on soft channels. But the initial development of such molds still requires considerable effort and expense.
∗
Corresponding author. Tel.: +886 5 2720411x66410; fax: +886 5 2721040. E-mail address: [email protected] (S.-C. Wang).
0021-9673/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2005.10.039
Recently, machining microchannel devices with simple industrial CO2 laser marker on plastic substrates such as PMMA have been reported for the applications of nucleic acid hybridization and optical sense using [5,6]. These laser markers have reasonable cost (20k US dollars or less) and are capable of easily fabricating typical two-dimensional micro-structures. Besides, no extra cost of either hardware or software development is necessary to fabricate microchannels at moderately accelerated throughput of approximately tens of chips per day. The production scale and initial cost fits the demand of a bio-analytical laboratory, wherein versatility is equally important as economical advantages in developing tens of chips for preliminary evaluation. To sustain adequate electroosmotic flows in microfluidic devices for fluid pumping, surface treatment on PMMA channel walls using strong reduction reagents has been studied [5]. The use of UV lasers to initiate photochemical reactions on plastics surface is an alternative protocol to generate electroosmotic flows in PMMA microchannel by charging the initially neutral acrylic substrate [7]. In contrast, generating electroosmotic flows in co-polyester microchannels is much easier. Ester functionality produces charged carboxylate group via hydrolysis when the channel is filled with diluted basic or acidic buffers [8,9]. The acidity dependence of electroosmotic mobility in co-polyester channels confirms the above mechanism [10]. Although
S.-C. Wang et al. / J. Chromatogr. A 1111 (2006) 252–257
co-polyester substrate absorbs UV and blue light considerably, this optical interference problem is curable with confocal detection, polarized fluorescence detection, or analyte velocity modulation method [11–14]. In this paper, we use a simple industrial CO2 laser marker to craft microchannels on Vivak co-polyester chips. Built-in pattern-marking software is used to control the writing motions of a low-power laser beam. To avoid overheating the plastic substrate, dry ice is placed in the marking chamber to significantly suppress the partial pressure of oxygen. Because the laser power is low, the laser beam has to move repetitively along the same line on a chip for several times to write one microchannel. The typical channel width is 300–350 m. The number of repetitive writing controls the aspect ratio of laser-ablated microchannel. We have measured the electroosmotic mobility in a copolyester microchannel after hydrolyzing it with buffers. The mobility is almost the same as that in the wire-imprinted microchannels reported previously [9,10]. The mobility is also stable whose measurement variation is less than 0.2%. Using surfactant cetyltrimethylammonium bromide (CTAB), we also confirm that dynamic coating protocols in silica capillaries are transferable to laser-marked plastic microchannels. Using video microscopy to measure the electrophoretic mobility of latex particles in the microchannel, we obtain the mobility histograms of particle mixture showing clear difference between bare and fluoroscein-tagged particles. 2. Experimental 2.1. Materials Co-polyester plastic slides are cut from Vivak plastic sheets (DSM, Sheffield, MA, USA). ACS grade reagents, including boric acid, sodium carbonate, acetic acid, citric acid, and sodium hydroxide, are used for the preparation of buffers. Cetyltrimethylammonium bromide is obtained from Sigma (St. Louis, MO, USA). We purchase bare (5 m) and fluorescein-tagged (10 m) latex particles from Fluka (Switzerland) and Polysciences (PA, USA), respectively.
253
the repetition count controls the aspect ratio of laser-ablated microchannel. To inspect the marked channel from the top view, optical microscopy is employed. The cross section of a channel is imaged using scanning emission microscopy (S-2400, Hitachi, Japan). 2.3. Measurement of electroosmotic mobility in microchannel We flush Tris–Borate–EDTA (TBE) buffer through our lasermarked microchannel to hydrolyze co-polyester substrate on the wall into the charged carboxylate functionalities. These charged moieties are able to establish an electrical double layer within the electrolyte solution. Electroosmotic flows are then produced when an electric field (50–400 V/cm) is applied across the channel. A current monitoring method is adopted to measure the current transition duration time during the replacement of a higher conductivity buffer with a lower one using electroosmosis [15]. Typically, dilute buffer is filled in the cathode reservoir to aspirate into the uncoated channel using electroosmosis after the electrophoresis voltage is turned on, and the dilute buffer gradually takes over the higher concentration buffer filled in the channel. Current through the channel is recorded with the voltage drop across a high resistor between the anode electrode and the ground. We employ four types of buffer solutions, sodium citrate, sodium acetate, sodium carbonate, and sodium borate, all at 25 mM to measure electroosmotic mobility at pH 3, 5, 7, and 9, respectively. Since the electroosmotic mobility is linearly proportional to the charge density on the channel surface (Helmholtz–Smoluchowski equation), the relative ratios of negative charge density on the channel surface between these pH values are obtained from the mobility data. The mobility values in CTAB-coated channels are also obtained using the current monitoring method at the same pH conditions as those described above. Since the surface of coated channel is positively charged, the direction of the electroosmotic flow is opposite to that in uncoated channel. Therefore, dilute buffer is filled in the anode reservoir to aspirate into the channel to replace higher concentration buffers.
2.2. Fabrication of microchannels We use an industrial CO2 laser marker (Model TL-25P, TopDog Laser, Taiwan) to fabricate microchannels on co-polyester chips. The laser beam has a wavelength of 10.6 m and the focus spot is approximately 100 m in diameter. The lasing tube is housed in a rack, whose motion is controlled by the buildin software. The housing rack is free to move on a horizontal surface in a chamber (70 cm × 60 cm × 26 cm) to strike the laser beam within an area of 24 cm × 24 cm. The writing speed of the laser beam is 15 cm/s. Approximately 100 g of dry ice is placed in the marking chamber to significantly dilute the partial pressure of oxygen. During the marking procedures, the laser power is intentionally reduced to 1.4 W to avoid overheating the chip. We control the laser beam to move repetitively along the same line on the chip for several times to machine one microchannel;
2.4. Observation of latex particle migration in microchannel to distinguish electrophoretic mobility We use a microscope to trace the movement of bare and fluorescein-tagged latex particles in the microchannel. Treating the microchannel surface with borate (25 mM) buffer, we apply an electric field of 75 V/cm across the channel filled with borate (25 mM). The concentrations of bare and functionalized latex particles seeded in the microchannel are 9.1 × 105 and 1.82 × 105 particles/ml, respectively. Although the particles are neutral or negatively charged, they move toward the ground electrode due to the electroosmotic flow. With a CCD camera (DFW-V500, Sony, Japan) operating at the acquisition rate of 15 frame/s, we acquired the images of particle movement under the focus region of an objective lens on the Nikon microscope.
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Particle trajectory and speed is analyzed with image processing software (ACDSee 6.0, ACD Systems, Canada). 3. Results and discussion Fig. 1A and B show the photographic images of the CO2 laser-marked channels on co-polyester slides using the proposed fabrication procedures. Fig. 1A shows a straight channel fabricated with the protocols described in Section 2. The serpent channels illustrated in Fig. 1B, on the other hand, can be fabricated by first machining the horizontal channels and later marking the connecting vertical channels with similar protocols. The widths of these channels are between 300 and 350 m. Therefore, the proposed procedures are demonstrated to be capable of marking straight or serpent channel. Fig. 2 shows the SEM picture of a cross-section of a laser-marked channel with onehalf aspect ratio. The channel surface can be observed to be adequately smooth. The SEM pictures (not shown) of channels at different aspect ratio (approximately 0.5–2.0) indicate similar smoothness.
Fig. 2. SEM image depicting the cross-section of a co-polyester microchannel created with CO2 laser marker. The channel aspect ratio is 0.53.
We used current monitoring method to measure electroosmotic mobility in the single straight channel chip (Fig. 1A). A typical curve showing the process of replacing the concentrated buffer (sodium acetate; 25 mM) with a dilute one (20 mM) is illustrated in Fig. 3. The current across the channel gradually drops while the dilute buffer is being filled into the channel. Except at the beginning and the end of the buffer replacement process, this current drop curve is almost linear. The regression result using the data of linear curve shows the correlation coefficient (r) is 0.997. The standard deviation of residue values (i.e. standard error) is 3.08 × 10−4 mA, which is less than 0.2% of the average current level (2.66 × 10−1 mA) of 25 and 20 mM buffer. Since the electroosmotic mobility variation during the buffer displacement process in the channel is virtually negligible, the mobility is therefore proved stable. Although the channels are not perfectly straight, slight irregularities in the channel do not influence the electroosmotic flow stability, as will be subsequently discussed. We also measure electroosmotic mobility in three different single channel chips (Fig. 1A) cut from the same
Fig. 1. (A) Single segment of a straight microchannel fabricated on a copolyester chip with CO2 laser marker. (B) Co-polyester substrate fabricated with a serpent channel using CO2 laser marker. The channel width in parts A and B is 324 m. The channel bottoms have been dyed with ink to improve visualization.
Fig. 3. The channel current monitored in the process of displacing 25 mM acetate buffer solution with 20 mM solution. The straight line shows the linear regressed curve using the data in the middle of displacement process.
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Fig. 4. The effect of buffer pH values on electroosmotic mobility in hydrolyzed and surfactant-coated co-polyester microchannels. The upper trace shows the buffer effect in a bare channel. The electroosmotic mobility in pH 3 is too small to detect. The lower trace shows the effect in a coated channel. The coating reagent is cetyltrimethylammonium bromide (CTAB).
plastic sheet. These mobility values were found to be 2.9 (±0.15) × 10−4 cm2 /V s in acetate buffer (pH 4.8). This negligible variation (less than 3.6%) in the measured mobility is consistent with the reported value in wire-imprinted channel [9,10]. We illustrate the pH dependence on electroosmotic mobility data measured with the current monitoring method in bare hydrolyzed co-polyester microchannels and surfactant-coated ones in Fig. 4. The curve for the bare channel keeps increasing as buffer acidity changes from acidic to neutral and becomes almost constant above pH 6. The other curve for the CTABcoated channel indicates the similar pH dependence of electroosmotic mobility to that in hydrolyzed co-polyester microchannels. Although the direction of electroosmotic flows in coated channels is reversed, the flow rate also increases from acidic to neutral pH conditions and become constant. The trends of these two curves are similar to those in previous reports using wire-imprinted channels [9,10]. Fig. 5 shows five consecutive images tracing the migration of bare latex particles seeded in the microchannel filled with sodium borate buffer (25 mM; pH 9). The positions of three traced particles driven in the electric field 75 V/cm are highlighted by circle, square, and oval marks. The outward arrows from the marks show the direction of the particle as it translates. Clearly, the migration of these particles is unidirectional suggesting the absence of any circulation in the bulk flow due to variations in the electroosmotic flow on the channel wall [16]. This observation indicates the negligible mobility variation in the channel, which is consistent with the conclusion obtained with current monitoring method in the above. Theoretically, the stability of electroosmotic flow rates is not sensitive to the variation of channel width when the charge density of the channel wall is homogenous [17]. Since the hydrolysis conversion of co-polyester to carboxylate takes place in the channel without spatial preference, the wall charge density distribution has to be uniform. When the characteristic length of channel is much larger than the Debye length, typically 10–100 nm in dilute electrolyte solutions, the Smoluchowski slip
Fig. 5. Five consecutive image snapshots taken once a second showing the migration trajectories of three latex particles moving across one 1500 m zone under the microscope objective. The traced particles are indicated with circle, square, and oval marks, respectively. The horizontal arrows show the migration direction of particles. The channel is filled with sodium borate buffer (25 mM; pH 9). The electric field applied across the channel is 75 V/cm.
velocity, which is proportional to the tangential field, applies at the slip plane. Since there is normal velocity on the surface, a continuous surface is then covered by the streamlines of the flow. As the surface charge screens the external field, the electric field is exactly tangential to the charged surface anywhere. Moreover, the governing equations for the interstitial flow driven by the slip velocity on the surface is the Laplace equation for the velocity potential, which is identical to the Laplace equation for electric potential. Such duality of boundary conditions and governing equations for the velocity potential and electric potential implies that the electric field lines and the streamlines are identical in the bulk if the surface charge is uniform. This has been proven by Takhistov et al. [18]. As a consequence of this duality, the flow rate is identical to the current for a channel with uniform charge. Another corollary is that, while the total flow rate is dependent on the transverse geometry and dimension of the channel, the electroosmotic mobility and local velocity are not. Our electroosmotic mobility measurement data in ablated co-polyester channel show almost equivalent mobility to that in fused-silica capillary. This result also suggests that the channel surface is smooth, not coarse. With uneven roughness, the local slip velocity would not be uniform and pressure-driven back flow would result producing vortices and transverse particle trajectories such as those observed by Minerick et al. [16] and Takhistov et al. [18]. Our data show that the unidirectional longitudinal particle migration driven by electroosmosis and the steadiness of buffer displacement process in the channel are both consistent with our theoretical conjecture that the channel is smooth.
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Although electro-osmotic mobility in the CO2 laser-ablated channel is steady, capillary electrophoresis (CE) separations in the channel is not feasible because the channel width (300–350 m) is not narrow enough. The plastic conduit fabricated with the laser marker is somewhat short of surface force to resist the hydrostatic pressure across the channel. The surface force is proportional to the channel width but the hydrostatic force is proportional to the channel cross-section (the square of channel width). When channel width increases, the hydrostatic force exceeds the surface force. Therefore, analyte loading
becomes questionable for sustaining limited dispersion of sample zone. Since the channel mobility is stable, particles with different electrophoretic mobility are distinguishable in the channel. Using particle trajectory data acquired with video microscopy, particle mobility values can be obtained. Fig. 6A and B show the individual mobility histograms of bare and fluorescein-tagged latex particles in the microchannel, respectively. The inset figures are the simulated electropherograms, whose peaks are depicted using the mean values and standard deviations obtained as shown in Fig. 6A and B. When the channel is filled with borate buffer (25 mM; pH 9.2), its electroosmotic mobility of 6.4 × 10−4 cm2 /V s is determined. The surface charge of bare latex particle is usually small. Therefore, the apparent mobility of bare particles in the hydrolyzed channel should be very close to the electroosmotic mobility of channel. The apparent mobility of bare particle on average is estimated as 6.4 × 10−4 cm2 /V s. This mobility value is reasonable. On the other hand, fluoresceintagged particles are more negatively charged. The mean apparent mobility of these functionalized particles should be somewhat less than the channel mobility, which is consistent with the result, 3.7 × 10-4 cm2 /V s, determined from the histogram of Fig. 6B. The Fig. 6C shows the mobility histogram of particle mixture in the same channel. This histogram has two distinct clusters, whose mean mobility values, 6.4 × 10−3 and 3.7 × 10−4 cm2 /V s, respectively, are identical to those estimated in Fig. 6A and B. The peaks in the simulated electrophoregrams in the inset figures are depicted using the mean values and standard deviations of two clusters in Fig. 6C. Having been easily classified as two distinct clusters, the mobility values of two bare and functionalized latex particles are distinguishable in the laser-ablated co-polyester channel. 4. Conclusions
Fig. 6. The electrophoretic mobility histograms in the channel of bare latex particles frame A, fluorescein-tagged latex particles frame B, and the mixtures of two types of latex particles frame C. The channel is filled with borate buffer solution (25 mM). The insets in A–C show the simulated electropherograms converted from the mobility histograms.
We have successfully developed a technique to fabricate microchannels on co-polyester substrates using an industrial CO2 laser marker. When the laser power is weak and partial pressure of oxygen in the machining chamber has been suppressed, repetitive writing motions of the laser beam on the chip gradually machine a microchannel of 300–350 m width. By controlling the writing power and repetition times, we are able to fabricate the microchannel at varied aspect ratios from approximately 0.5–2. Similar to wire-imprinted channels, laser-ablated copolyester channels undergo hydrolysis to become charged. Although the channel width is not perfectly consistent, stable electroosmotic mobility is validated in the laser-ablated channel. In addition, we show that dynamic coating protocols in capillary electrophoresis are transferable to the laser-marked channel. This channel width is too wide to provide adequate surface force to avoid solute dispersion in the sample loading process for performing capillary electrophoresis separations. By using video microscopy we demonstrate that CO2 laser-ablated channel is suitable to distinguish bare latex particles from functionalized ones with the particle mobility histograms.
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Acknowledgements The authors express thanks for the financial support from National Science Council, Taiwan (NSC92-2113-M-194-022 and NSC93-2120-M-194-005), and National Chung Cheng University. The authors also acknowledge helpful discussions with L.Y. Yeo and H.-C. Chang (Department of Chemical and Biomolecular Engineering, University of Notre Dame, IN, USA) in the preparation of this manuscript. We also thank J.-Y. Cheng (Center for Applied Sciences, Academia Sinica, Taiwan) for discussions on laser-marking techniques. References [1] T.D. Boone, Z.H. Fan, H.H. Hooper, A.J. Ricco, H. Tan, S.J. Williams, Anal. Chem. 74 (2002) 78A. [2] A.Y. Fu, C. Spence, A. Scherer, F.H. Arnold, S.R. Quake, Nat. Biotechnol. 17 (1999) 1109. [3] M.A. Witek, S.Y. Wei, B. Vaidya, A.A. Adams, L. Zhu, W. Stryjewski, R.L. McCarley, S.A. Soper, Lab on a Chip 4 (2004) 464.
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