Trapping of Bioparticles via Microvortices in a Microfluidic Device for

Oct 28, 2008 - Georgette B. Salieb-Beugelaar , Giuseppina Simone , Arun Arora , Anja Philippi and Andreas Manz. Analytical Chemistry 2010 82 (12), 484...
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Anal. Chem. 2008, 80, 8937–8945

Trapping of Bioparticles via Microvortices in a Microfluidic Device for Bioassay Applications Cheng Ming Lin, Yu Shang Lai, Hsin Ping Liu, Chang Yu Chen, and Andrew M. Wo* Institute of Applied Mechanics, National Taiwan University, Taipei, Taiwan This paper presents hydrodynamic trapping of bioparticles in a microfluidic device. An in-plane oscillatory microplate, driven via Lorentz law, generates two counterrotating microvortices, trapping the bioparticles within the confines of the microvortices. The force required to trap bioparticles is quantified by tuning the background flow and the microplate’s excitation voltage. Trapping and releasing of 10-µm polystyrene beads, human embryonic kidney (HEK) cells, red blood cells (RBCs), and IgG antibodies were demonstrated. Results show the microvortices rotates at 0-6 Hz corresponding to 2-9 Vpp (peak-to-peak) excitation. At a particular rate of rotation (2-7 Vpp tested), a bioparticle is trapped until the background flow exceeds a limit. This flow limit increases with the rate of rotation, which defines the trap/release force boundary over the range of operation. This boundary is 12 ( 2.0 pN for cell-size bioparticles and 160 ( 50 fN for antibodies. Trapping of RBCs demonstrated microvortices’ ability for nonspherical cells. Cell viability was studied via HEK cells that were trapped for 30 min and shown to be viable. This hydrodynamically controlled approach to trap a wide range of bioparticles should be useful as a microfluidic device for cellular and subcellular bioassay applications. The lab-on-a-chip approach leverages upon microfluidics technology1 and is becoming an enabling platform for miniaturization of biological and chemical analyses. Characteristically, these devices require small sample volume, fabricate with relative ease, are conveniently controlled,2 and often utilize rapid detection process.3 They provide a suitable environment for studies of biosamples while integrating various analytical operations in a perfusion system.4 For example, microfluidic system is capable of real-time bioassays of controlling fluid conditions, including exchange of fluid media,5 supply of sufficient grow factors, and refresh fluid for live cells and biomolecules.6,7 In cellular studies via microfluidics, some form of cellpositioning method is often needed. Invasive (contact) cell * To whom correspondence should be addressed. E-mail: andrew@ iam.ntu.edu.tw. (1) Andersson, H.; vanerg, A. Sens. Actuators, B: Chem. 2003, 92 (3), 315– 325. (2) Rhee, S. W.; Taylor, A. M.; Tu, C. H.; Cribbs, D. H.; Cotman, C. W.; Jeon, N. L. Lab Chip 2005, 5 (1), 102–107. (3) Cheng, X.; Liu, Y. S.; Irimia, D.; Demirci, U.; Yang, L. J.; Zamir, L.; Rodriguez, W. R.; Toner, M.; Bashir, R. Lab Chip 2007, 7 (6), 746–755. (4) Easley, C. J.; Karlinsey, J. M.; Landers, J. P. Lab Chip 2006, 6 (5), 601– 610. 10.1021/ac800972t CCC: $40.75  2008 American Chemical Society Published on Web 10/29/2008

trapping can provide direct control but faces challenges due to the complex physical properties of biosamples; for instance, white blood cells are very sticky while RBCs are rather nonadhesive. Carlson et al. have designed mechanical filters for trapping different cell types due to their different physical properties.8 Moreover, noncellular entities with similar properties to cells were trapped at a fixed location using micropipet-like approaches,9,10 special microstructure aides with hydrodynamic force.11 However, among other undesirable effects, direct physical contact approaches often cause contamination to biosamples and result in one-time usage of the device. A noninvasive manipulation method to control cells is preferable. An ideal cell diagnostic platform should retain the natural properties of cells and, if needed, provide a highly efficient medium exchange with limited side effects to cells. Such medium exchange can supply nutrients and remove metabolites, along with ensuring a stable and gentle environment for live cells.12 Cell manipulating using a noninvasive approach is the best approach to handle nonadherent cells for bioassays.13 Various noninvasive cell trapping techniques have been used, e.g., optical tweezers (OT),14 dielectrophoretic force,15 acoustics,13,16 and a hydrodynamic method.17 Optical tweezers use a focused laser beam to generate forces on a cell based on the difference in the refractive index between the cell and the medium. Trapping is achieved by focusing the laser beam through the microscope objective. Sylvie Henon et al.18 used the OT to hold a red blood (5) VanDelinder, V.; Groisman, A. Anal. Chem. 2007, 79 (5), 2023–2030. (6) Lu, H.; Koo, L. Y.; Wang, W. C. M.; Lauffenburger, D. A.; Griffith, L. G.; Jensen, K. F. Anal. Chem. 2004, 76 (18), 5257–5264. (7) Albrecht, D. R.; Underhill, G. H.; Mendelson, A.; Bhatia, S. N. Lab Chip 2007, 7 (6), 702–709. (8) Carlson, R.; Gabel, C. V.; Chan, S. S.; Austin, R. H.; Brody, J. P.; Winkleman, J. Phys. Rev. Lett. 1997, 8, 2407–2407. (9) Pantoja, R.; Nagarah, J. M.; Starace, D. M.; Melosh, N. A.; Blunck, R.; Bezanilla, F.; Heath, J. R. Biosens. Bioelectron. 2004, 20 (3), 509–517. (10) Chen, C. Y.; Liu, K. T.; Jong, D. S.; Wo, A. M. Appl. Phys. Lett. . 2007, 91, 12. (11) DiCarlo, D.; Aghdam, N.; Lee, L. P. Anal. Chem. 2006, 78 (14), 4925– 4930. (12) Powers, M. J.; Janigian, D. M.; Wack, K. E.; Baker, C. S.; Stolz, D. B.; Griffith, L. G. Tissue Eng. 2002, 8 (3), 499–513. (13) Evander, M.; Johansson, L.; Lilliehorn, T.; Piskur, J.; Lindvall, M.; Johansson, S.; Almqvist, M.; Laurell, T.; Nilsson, J. Anal. Chem. 2007, 79 (7), 2984– 2991. (14) Ashkin, A. Biophys. J. 1992, 61 (2), 569–582. (15) Voldman, J.; Gray, M. L.; Toner, M.; Schmidt, M. A. Anal. Chem. 2002, 74 (16), 3984–3990. (16) Wiklund, M.; Toivonen, J.; Tirri, M.; Hanninen, P.; Hertz, H. M. J. Appl. Phys. 2004, 96 (2), 1242–1248. (17) Shelby, J. P.; Chiu, D. T. Lab Chip 2004, 4 (3), 168–170. (18) Henon, S.; Lenormand, G.; Richert, A.; Gallet, F. Biophys. J. 1999, 76 (2), 1145–1151.

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cell (RBC) and determined the shear modulus of cellular membrane. Dielectrophoresis (DEP) can trap polarizable bioparticles under a nonuniform electric field. The approach has been well studied due to its relative ease of fabricating planar electrodes in a myriad of configurations, each resulting in a unique electric field for a specified task. Voldman et al.15 have shown that a negative DEP configuration induced effective dipole moment in a cell that is antiparallel to the electric field, which creates a dielectrophoretic force that could trap the cell stably. Also, Becker et al.19 showed that breast cancer cells could be attracted to electrode tips via positive DEP traps, suggesting proper design of electrodes can trap and move the cell to a predetermined position. However, polarization of cells induced by electric field may affect their interior. Perhaps the most severe limiting factor for utilization of DEP is the need to employ suitable combination of medium and cell, based on their dielectric (not physiological) properties, in order to produce a desired manipulative force. Another noncontact method is acoustical tweezers, which generates a trapping force via the difference in compressibility between a cell and the medium. The induced pressure difference aggregates the cells by controlling the acoustical energy. This technique usually uses an ultrasonic standing wave to trap particles and cells.20,21 It provides a rapid and reliable trapping quality for cells and particles.13,20 Ultrasound has been used for many years in the studies of microscale particles. By changing operation frequency and reflecting distance, the trapping forces can be controlled and specific particle sizes can be trapped. There have been several noninvasive hydrodynamic-based studies to trap cells in recent years, with one of the main differences in the method of generating the needed trapping mechanism. Shelby et al.22 have utilized microvortices to manipulate bioparticles and measured their rotational rate within a confined region, and application of the technique on nanoparticles has been demonstrated.17 The flow in which cells were trapped was induced in a novel manner by the main channel (steady) flowsthrough an opening in the channel wallsleading to secondary flow in the cavity where cells were located. Lutz et al.23 were the first to utilize secondary streaming flow to trap cells behind a fixed cylinder and measured the motility force of motile cells. The streaming flow in this hydrodynamic tweezers23 approach was generated by an external piezoelectric apparatus. Hence, one can quantify the trapping force by varying the frequency or amplitude of the streaming flow. Although these excellent works have provided progress on hydrodynamic trapping, much work is still needed in exploiting the full potential of the advantages of the method, e.g., providing a gentle environment for biosamples. In this study, hydrodynamic trapping of bioparticles was demonstrated in a microfluidic device utilizing a resonating microplate driven by Lorentz force, generating two counterrotating microvortices. After quantifying the characteristics of the (19) Becker, F. F.; Wang, X. B.; Huang, Y.; Pethig, R.; Vykoukal, J.; Gascoyne, P. R. C. Proc. Natl. Acad. Sci. U. S. A. 1995, 92 (3), 860–864. (20) Hertz, H. M. J. Appl. Phys. 1995, 78 (8), 4845–4849. (21) Bazou, D.; Kuznetsova, L. A.; Coakley, W. T. Ultrasound Med. Biol. 2005, 31 (3), 423–430. (22) Shelby, J. P.; Mutch, S. A.; Chiu, D. T. Anal. Chem. 2004, 76 (9), 2492– 2497. (23) Lutz, B. R.; Chen, J.; Schwartz, D. T. Anal. Chem. 2006, 78 (15), 5429– 5435.

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Figure 1. Illustration of flow field behind a backward facing step with a separation vortex, which can be used to trap particles within the recirculating flow region.

microvortices, studies were conducted on the force required to trap/release bioparticles, which include 10-µm polystyrene beads, human embryonic kidney (HEK) cells, RBCs, and IgG antibodies. A cell viability study was also undertaken to scrutinize the cell condition after being trapped to ensure utilization of the microdevice. DESIGN OF THE DEVICE Overall Concept. The force acting on a suspended bioparticle under some local flow condition is proportional to the fluid properties, the bioparticles’ geometry, and the velocity difference between the bioparticle and the flow. If a desirable local flow can be generated, the resultant hydrodynamic force might be suitable for manipulating bioparticles, avoiding the undesirable effects of physical contact methods. In mesoscale flow, a vortex is often generated via relatively high speed flow over a sharp corner or a cavity. As such, a suspended particle could enter the vortex and be trapped within the recirculation zone, as illustrated in Figure 1. In microscale, however, most microdevices operate in low Reynolds number, posing generation of vortices a difficult task, much less utilizing them to trap bioparticles. Design of Microvortices Generator. Lorentz force was utilized to drive a microplate to resonance (140 kHz). One advantage of using Lorentz force is to reduce the complexity of actuation of the MEMS-based device by employing an external magnet. Once in resonance, the microplate generates two microvortices near its edges, providing a local flow condition for hydrodynamic manipulation of bioparticles. The main structure of the vortices generator consists of a double-clammed, suspended bridge, combined with a square plate in the middle as the primary structure, as shown in the 3D sketch of Figure 2a. When an alternating current (ac) passed through the gold layer on the surface of the suspended bridge in the presence of an external magnetic field B (Nd-Fe-B magnet, ∼1 T) perpendicular to the bridge surface, the main structure was forced to resonate in the third (in-plane) direction. The geometry of the bridge is 1.2 µm thick, 20 µm wide, and 750 µm long, with the square microplate (100 µm by 100 µm) in the middle. This design is distinct from that of Lutz et al.23 in several ways. First, the method of generation is entirely different: the driving source is embedded within the present device versus external.23 Second, the region where cells are trapped is in the outer streaming flow in this study versus within the inner eddies. Third,

Figure 2. Sketch of the device and the fabrication process. (a) Design of the device showing a suspended structure with a square plate in the middle. Structure was driven to resonance laterally (inplane) at 140 kHz via Lorentz lawsforce induced from an ac current passes through the gold layer on the structure in the presence of an external magnetic field B. The gap between suspended structure and the silicon nitride is 20 µm. (b) Fabrication processes. First, the silicon substrate was deposited on silicon nitride (∼1 µm) (step 1). Then, metallic electrodes (0.2-µm thickness of Au/Cr) were sputtered on silicon nitride (step 2). Then, electrodes were patterned using photolithography with photoresist (PR) and metal etchant (step 3). By repeating the processes, the window of the cavity underneath oscillatory structure was formed (steps 3 and 4). The silicon nitride unprotected by photoresist would be etched using reactive ion etching (RIE) (step 5). Finally, the device was etched utilizing potassium hydroxide etchant (step 6).

the frequency of oscillation was fixed at 140 kHz versus 40 Hz to 1 kHz. These differences have consequences in the entire flow field and, more importantly, on the trapped bioparticles, e.g., shear stress, as will be discussed below. Quality Factor. This vortices generator is a resonant-based actuator operating under liquid with the natural frequency ∼140 kHz. This is implied from the rotational velocity of the vortices, which maximizes at this frequency and decays substantially at off-resonance frequencies. The maximum displacement of the oscillatory plate is less than 1 µm. From these characteristics, the quality factor (Q) is calculated to be ∼10, which is indicative of a reasonable resonating device. EXPERIMENTAL ASPECTS Microfabrication. Fabrication of the suspended structure utilized conventional lithographic microfabrication, as shown in Figure 2b. First, silicon nitride (∼1 µm) was deposited on the silicon substrate with a low-pressure chemical vapor deposition system. Then, a metallic layer (0.2-µm thickness of Au/Cr) was sputtered on the silicon nitride surface. A photoresist (S1813, Shipley) layer defined the microelectrode structure. Unnecessary metal was removed by metal etchant. Next, the positive photoresist

on the metal is removed by acetone. By repeating the procedure of coating photoresist and development, the open window of the cavity underneath the oscillating structure was formed. The silicon nitride not covered with the photoresist would be etched using RIE. Then, a bulk micromachine process was used to remove unwanted silicon below the plate/beam structure using potassium hydroxide, thus suspending the structure. Only two masks are needed in the fabrication procedure. The microchannel was fabricated with poly(dimethylsilane) (PDMS, Sylgard 184, Dow Corning) via a soft lithographic technique.24 The ratio of elastomer/curing agent used was 10:1 by weight, mixed uniformly and degassed at low pressure for 20 min. Next, the premixed PDMS was poured on a master and baked at 85 °C for 1 h. After cutting and peeling off from the master, the replica PDMS channel is formed. There are two different PDMS channel height in this work: (1) 1000 µm for characterizing microvortex and trapping force (used in Figures 5, 7, and 8 and (2) 50 µm for RBCs and cell viability test (Figure 9). Device Assembly. The integrated microfluidic device is composed of a silicon chip (28 × 20 × 0.5 mm3) that houses the vortices generators, a PDMS microchannel, and a poly(methyl methacrylate) (PMMA) board (50 × 50 × 3 mm3), as shown in Figure 3. A linear array of four vortex generators in a silicon chip was mounted on PMMA for easy handling. A PDMS channel was bonded to the surface of the silicon chip using an oxygen plasma treatment. Polyethylene tubings (1.09-mm o.d. and 0.38-mm o.d.) were connected to fluid connection components on the back side of PMMA, providing a convenient method for cell and medium injection by a syringe pump. PMMA board includes three drilled holes: two inlet holes and one outlet. A permanent magnet was placed directly beneath the oscillators, with it large enough that the magnetic field can be considered uniform for all oscillators. Procedures. The chip and other parts of the microfluidic device were first sterilized using 70% ethanol. Then, bioparticless polystyrene beads, cells, and antibodiesswere injected into the device to study the trapping characteristics of the microvortices. Further details are provided in subsequent sections. For trapping of cell-sized bioparticles (10 µm), an optical microscope was used to observe the process of vortex generation and to study the trapping/releasing phenomenon. Additionally, a CCD camera (Unibrain, Fire-i400) recorded images for postprocessing, at the maximum frame rate of 30 frames/s at resolution of 640 × 480. The interface IEEE1394 provides sufficient bandwidth for data transmission from CCD to PC. To probe the trapping performance of bioparticles smaller than cellular level, biomolecules with the effective Stokes diameter of 10 nm were studied. The trapping experiment was done via the same device and measurement procedure but the operation was observed with an inverted fluorescent microscope (Olympus IX71) with a CCD camera (Olympus DP-70). Bioparticles and Preparation. Biological particles vary greatly in dimensions and shapes, which might affect the effectiveness of their trapping via the microvortices. Hence, a range of bioparticles were studied to scrutinize the generality of usage of the device. Bioparticles tested included the following: 10-µm polystyrene beads, human embryonic kidney (HEK-297T, ATCC) (24) Xia, Y. N.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37 (5), 551– 575.

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Figure 3. Microdevice. (a) Top view showing four oscillators as microvortices generators integrated in a silicon chip (28 × 20 mm2) bonded between a PDMS channel and PMMA substrate (50 × 50 mm2). Two wires were clipped to the metallic pad for electric connection to the oscillators. (b) Side view sketch of the device illustrating the overall structure and arrangement.

cells, RBCs, and goat anti-rabbit IgG (Alexa Fluor 546, Molecular Probes, Eugene, OR, Catalog No. A-11010). The cell-size polystyrene beads analogically performed as cells trapped by the microvortices for the initial observation. The HEK cells represented behavior of living cells within the hydrodynamic environment of the microvortices and allowed testing of cell viability. The RBCs are nonspherical in shape and thus serve to characterize the efficacy of the microvortices in regard to shape dependency. Finally, the use of much smaller (∼10 nm) biomolecules would test the extent of generality of trapping/releasing claim of the device. Details of preparation of HEK cells and RBCs are provided as follows. HEK cells were grown in Dulbecco’s modified Eagle medium (Gibco/Invitrogen) supplemented with 10% fetal bovine serum (Gibco/Invitrogen). Then, 1% penicillin-streptomycin and 1% sodium pyruvate were added at 37 °C in a humidified environment of 5% CO2 in air. Prior to injecting into the microdevice, cells were detached from the culture dish, treated in trypsinethylenedinitriletetraacetic acid (trypsin-EDTA) in a 37 °C incubator for 4 min, and then resuspended in medium. Cells and medium were then centrifuged at 1000 rpm for 5 min. The supernatant was aspirated, and cells were resuspended in 10 mL of PBS solution with a pH of 7.4. RBCs were separated from whole blood cells by centrifugation and, to alleviate clogging, diluted with PBS, achieving a density of 104 cells/µL. Cell viability test was conducted to ensure the microvortices platform is indeed appropriate for cellular studies. Acetomethoxy derivate of calcein (calcein AM, C3100MP, Invitrogen) was prepared in stock solution (50 µg of calcein AM in 9.6 µL of DMSO) and diluted with standard phosphate-buffered saline solution (PBS) in a volume ratio of 250. After HEK cells were trapped for 30 min, calcein AM was perfused into the channel, testing the cells for viability while cells were still rotating within the microvortices. Calcein AM can transport through the cellular membrane into cells then be hydrolyzed by an intracellular enzyme resulting in strong green fluorescence. As dead cells lack this enzyme, only live cells are marked. Fluorescent images were taken via the CCD camera (DP-70, Olympus) using an inverted microscope (IX-71, Olympus). Temperature Measurements. Generation of the microvortices requires an electric current flowing through the device, which results in fluid temperature increase. This temperature increase in microchannel can be considered theoretically as power dissipation from the oscillator surface when a current passing through a material, i.e., P ) V2/R, where I and V are the rootmean-square values of current and ac voltage, respectively. 8940

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Exposure of cells to high temperature would cause extra stress and results in irreversible damage. To probe the effect of temperature increase with various driving voltages, the fluid temperature within the microchannel was quantified at two typical background flow velocity, i.e., 20 and 100 µm/s. Characterization of temperature effect was conducted as follows. Temperature-dependent fluorescent dye, Rhodamine B (Sigma-Aldrich, Inc.), was used since its response is sufficiently rapid (ms) and with good spatial resolution (µm).25 With this technique, fluid temperature can be evaluated via fluorescent intensity, which would decrease as temperature increases. A solution of 0.1 mM Rhodamine B was injected into the microdevice and imagine acquisition was performed by a CCD camera (DP70, Olympus) mounted on an inverted fluorescent microscope (IX 71 Olympus, Hg lamp, 10× objectives). To calibrate the intensity with temperature, a thermal couple was placed in contact with the back side of the silicon chip bonded with PDMS channel. A heater was embedded in a Petri dish in which our device was placed upside down lying on the bottom of the dish. Then, the dish was filled with water in order to provide a stable temperature reservoir as the heater was activated. Fluorescent imagine of stable temperature from 25 to 70 °C was recorded. The calibration curve was generated by averaging the gray value of intensity corresponding to each temperature. By infusing the fluorescent dye into the activated microdevice, we can evaluate the temperature in the vicinity of the microvortices. Four sets of data were taken at each driving voltage. The gray value at room temperature was subtracted from the mean gray value of the intensity data at each temperature in order to reduce the environmental variations over time, such as lamp intensity. The standard variation of the temperature increase was less than 1 °C. The experiments were conducted for both 20 and 100 µm/s background flow velocity. Hydrodynamic Force. In this work, bioparticles were hydrodynamically trapped via the microvortices. Procedurally (see Figure 7a), bioparticles were pumped into the device upstream of the vicinity of the microvortices, with them rotating relatively rapidly. This ensures the bioparticles are trapped by the microvortices and rotate with the vortices while the background fluid passes by. Then, the background flow velocity was reduced, along with decrease in the excitation voltage until a particular voltage to be tested was reached. With the bioparticles still trapped and rotating with the microvortices, the flow was gradually increased until they were released. The entire process was recorded by CCD camera for postprocessing. The procedure was repeated for other (25) Ross, D.; Gaitan, M.; Locascio, L. E. Anal. Chem. 2001, 73 (17), 4117– 4123.

Figure 6. Temperature increase in fluid relative to room temperature (25 °C) vs voltage at two flow velocities. Voltage between 2 and 7 Vpp was used for trapping of bioparticles.

Figure 4. Counter-rotating microvortices. (a) Micrographs, taken from snapshots of movie frames (using a portable digital microscope, with a frame rate of 15 frames/s), showing time-lapsed trajectories of microvortices, made visible by trapped 10-µm polystyrene beads. The right vortex is shown to trap more particles than that of the left. Background flow, when applied, was from the left, along the plane of the microvortices. The device operated at a frequency of 140 kHz, with the microdevice placed in a large container filled with water. (b) Sketch of the trajectories of the microvortices above the edges of the oscillatory plate, as observed from Figure 4a. The orbits of the microvortices resemble a slightly inclined ellipse, with the major axis ∼100 µm. A movie is available in the Supporting Information.

Figure 5. Rotational velocity of a microvortex verses driving voltage (2-9 Vpp), with corresponding rotational frequency shown on the right ordinate. Results indicated the rotational velocity increases parabolically with voltage. The microvortex is very controllable and robust. Statistic analysis performed using Student’s t tests at 95% confidence level, n ) 3, S ) 1.0. The regression coefficient is 0.97.

excitation voltages. The device operated at a frequency of 140 kHz and voltage of 2-7 Vpp. The trapping force of the microvorticessa measure of the ability of a microvortex to confine a bioparticle rotating along its

circulatory streamlinesswas quantified. If the hydrodynamic force from the background flow acting on a bioparticle exceeds the maximum trapping force corresponding to a particular vortex rotational velocity, the bioparticle would be released from the microvortex, and conversely, a force from the background flow less than the maximum trapping force would ensure that the bioparticle remains in the trapped state. Thus, this maximum trapping force can calculated from the classical Stoke’s law for an object immersed in low Reynolds number flow, i.e., Fmax ) 6πηUa

(1)

where η is the viscosity, U is the velocity of the background flow that would cause release of a trapped bioparticle, and a is the radius of the bioparticle. RESULTS AND DISCUSSION Microvortices. The primary result of the in-plane resonating motion of the microplate is generation of two counter-rotating microvortices. Figure 4a presents a micrograph of the microvortices, showing time-lapsed trajectory of microvortices, made visible by trapped 10-µm polystyrene beads. Figure 4b sketches the trajectories based on postprocessing of dynamic image data. The trajectory indicates the flow within the vortex approaches the oscillatory plate from above, moves outward along the plate for a short distance, then leaves the plate area and curves upward, and eventually progresses downward toward the plate, completing the recirculating flow circuit. Once trapped, particles recirculate continuously within the confine of the microvortices just above the two edges of the oscillatory plate. The trapped particles are situated in a near-circular orbit (minor/major axes ratio of ∼4:5) concentrated along the outer rim. The width of orbit appears to depend on the uniformity of the trapped particles: 1-2 diameters for 10-µm polystyrene beads and 2-3 cell diameters for, say, HEK cells. However, the number of bioparticles entering each microvortex might not be the same, which is the case in Figure 4a. The microvortices are reliable and robust. The diameter of the microvortices ranges from 80 to 100 µm. Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

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Effect of Particle Size. Intuitively, particle size should affect the efficacy of the microvortices in confining the particles within the region of the recirculating flow since trapped bioparticles need to faithfully follow the elliptical streamlines of the microvortices against the possibility of being washed downstream by the flowing medium. Thus, a smaller particle should be more readily trapped than a larger one. To test this hypothesis, different sizes of polystyrene beadss1, 5, 10, and 15 µm in diameterswere tested in the microfluidic device. Trapping was commenced when suspended particles of a particular size flowed near the trapping zone and into the microvortices above the oscillatory plate. Results show all beads could be trapped under a variety of conditions, depending on the driving voltage and background flow. However, the 15-µm beads can escape from trapping easier than that of smaller particles. The 1-µm particles are sufficiently small that are very well confined within the microvortices. This size independency characteristic suggests that the microvortices might have general application as a trapping tool. Rotational Velocity of Microvortices. Rotational velocity of the microvortices should play a key role in defining the overall trapping characteristics since trapped particles are confined within the region of the microvortices, advecting along in elliptical orbit. Dimensionally, one expects rotational velocity depends on the oscillatory frequency of the microplate and a length scales microplate dimension, viscous penetration depth, or amplitude of oscillation. The data will provide clues to which length scale is correct. Figure 5 presents the results of rotational velocity for a range of driving voltage of the microplate. The average rotational velocity is in the order of hundreds to 1000 µm/s, which is a relatively fast value in microflow environment, and varies quadratically with driving voltage, having a threshold voltage of 2 Vpp. The second ordinate shows the rotational frequency of the microvortices, with maximum frequency of ∼6 Hz at 9 Vpp. Much smaller antibodies (10 nm) were also used as tracer particles in the rotational velocity experiment. Results showed the relationship between rotational velocity and driving voltage to be essentially equivalent to that of using polystyrene beads. In this work, with the geometry of the microplate and the frequency fixed, the variation of rotational velocity with voltage is largely due to variation of oscillatory amplitude with voltage (also see Schlichting28). It is interesting to note that the theoretical magnitude of the rotational (streaming) velocity does not depend on viscosity. Schlichting’s analysis of successive approximation28 (his eq 15.63) shows the streaming flow u2 far from the surface is u2(x, ∞) ) -

3 U0 dU0 4 n dx

(2)

where U0 is the velocity amplitude outside of the viscous region, n the radian frequency, and x the direction along the moving surface. This equation does not contain the viscosity explicitly. Experimentally, however, the value of U0 would depend on the viscosity implicitly since (as a reviewer pointed out) the viscosity would change by ∼15% due to temperature change. Viscosity affects the rotational velocity since it is a parameter in the timeaveraged governing equation for the streaming flow. Physically, as the driving voltage increases, both the Lorentz force and the fluid temperature increase. This results in two effects: (1) the 8942

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Figure 7. Trapping force measurement of cellular-scale microparticles. (a) Micrograph (top view) showing trajectories of particles entering, being trapped (black spots), and leaving the microvortices when the hydrodynamic force exerted by the background flow on the trapped particles is greater than the trapping force of the microvortices. (b) Trapping force of microvortices on 10-µm polystyrene beads under a range of parameters. The x-axis is the Reynolds number of the counter-rotating microvortices, expressed as ReΓ ) Γ/η, where Γ ) IV · dl, the circulation of a vortex. The operating voltage for trapping force measuring is from 2 to 7 Vpp. The Reynolds number scale (Ref ) U · h/η) on the far right ordinate characterizes the flow in the microchannel. The gray region represents the boundary between trapped and untrapped regions. Mean value and error bar range were calculated using Student’s t tests with 95% confidence level.

increase in Lorentz force causes an increase in the plate’s amplitude, and (2) the increase in temperature lowers the viscosity, which would further increase the amplitude due to decrease in viscous damping. Hence, both effects would contribute to the rotational velocity versus voltage data. Temperature Increases. The measured temperature increase by Joule heating showed the quadratic relation to drive voltage as we expected (see Figure 6.). Here, temperature rise is defined as the increase from 0 V condition, which is essentially at room temperature (25 °C). Two regression models of second order express the temperature response of fluid relative to room temperature under two different flow velocities in a microchannel. At drive voltage of 3 Vpp, the temperature increase is under 5 °C and the trapping force is good at 10-µm particles when the supplied background flow velocity was at 20 µm/s, producing an absolute temperature of 30 °C in the microchannel when room temperature was at 25 °C. With increasing background flow velocity to 100 µm/s, the temperature rise was only 2 °C. This temperature increase at a flow of 100 µm is less than that at 20 µm/s flow. Note that trapping beyond 7 V was not commenced; i.e., trapping data (in Figure 7) correspond to 2-7 Vpp only. This is because the temperature within the microdevice at 8V is beyond physiological temperature of 37 °C. However, measurement of rotational velocity was performed up to 9 Vpp. Trapping Force. Figure 7 presents the trapping force of the microvortices on cellular-sized particlessrepresented by measurement of 10-µm polystyrene beadssover a range of parameters. The trapping force is determined at the point that a vortex at a

particular rotational velocity can no longer retain a trapped particle against a particular background velocity, thus releasing it downstream. As shown in Figure 7a, this trapping force was quantified via a three-step process: first, observing particles that are embarking upon the trajectory to enter the microvortices; second, trapping particles at a particular vortex rotational velocity, i.e., at a particular driving voltage; third, releasing trapped particles by increasing the background flow velocity, while keeping the voltage fixed, until the particles leave the microvortices and progress downstream along the microchannel. This process was done with single or multiple particles (1–3, but more can be trapped). The point of release usually happens at the top of the microvortex far from the plate. By recording images of this process by a CCD camera (30 frames/s), the background flow velocity at which release occurs is fairly easy to determine. Figure 7b presents the results of the trapping force. The abscissa represents the Reynolds number of the microvortices, expressed as ReΓ ) Γ/η, where η is the kinematic viscosity and b is the circulation of the vortex at mean rotational Γ)Ib V · dL velocity (V) (from Figure 5) along the recirculation path (L), or ∼300 µm for 100-µm-diameter microvortices. Thus, the circulation corresponds directly with the driving voltage (see Figure 5). The far right ordinate characterizes the flow in the microchannel, represented by the Reynolds number Ref = U · h/η, where U is the background flow velocity and h the microchannel height. Results show a distinct trend of an increase in trapping force with increasing Reynolds number of microvortices, with the force level in pico-Newton range. The gray area corresponds to the maximum trapping force, or Fmax in eq 1, and divides the results into two regions, trapped or untrapped, and can be understood as follows. At a particular Reynolds number of microvortices, a particle is well trapped under low Reynolds number of background flow condition (below the gray area); hence the trapping force is sufficient to retain the particle within the trapped region. As the background flow increases (above the gray area), the particle escapes the microvortices, entering the untrapped region. Alternatively, at a particular background flow, a particle is not trapped at low microvortex rotational velocity (left of the gray area) and becomes trapped when the microvortex velocity is increased (right of the gray area). Thus, at a specified Reynolds number of background flow (Ref), particles can be trapped at a range of Reynolds number of microvortices (ReΓ) above the value defined by the gray area corresponding to that value of Ref. The maximum trapping force measured was found to be 12 ± 2.0 pN at the flow velocity of 140 µm/s and driving voltage of 7 V (peak-to-peak). It should be instructive to compare this force level with other noncontact methods. The cell-trapping force generated by optical tweezers is ∼100 pN,14,26 dielectrophoretic force in the range of 200-500 pN27 and acoustics 300-600 pN.13 The hydrodynamic tweezers23 can provide trapping force up to 30 pN, and the shear stress on cells is stated to be comparable to arterial shear stress in circulatory system, which is ∼1.5 N/m2. These above forces are that reported in the cited literature and should be capable of (26) Qian, F.; Ermilov, S.; Murdock, D.; Brownell, W. E.; Anvari, B. Rev. Sci. Instrum. 2004, 75 (9), 2937–2942. (27) Taff, B. M.; Voldman, J. Anal. Chem. 2005, 77 (24), 7976–7983. (28) Schlichting, H. Boundary-Layer Theory, 7th ed.; McGraw-Hill: New York, 1979; p 430.

Figure 8. Trapping force measurement of nanoscale bioparticles. (a) Micrograph trajectories of antibodies (IgG ∼10 nm) entering the microvortices (left dotted line) and upon release (right dotted line) when the flow of the PBS medium is excessive. Enrichment of antibody is demonstrated via increased fluorescence intensity as more antibodies are being trapped. Black dashed lines show the suspended plate. (b) The approximate trapping force exerting on the antibodies by the microvortices. Axes are similar to that of Figure 7. Results show the trapping force is in femto-Newton range. The operating voltage for measuring was from 2 to 7 Vpp.

even lower force level by a decreased voltage or optical power (as pointed out by one reviewer). Although the present approach is similar to the work on hydrodynamic tweezers23 in that the use of microvortices here for trapping is a form of hydrodynamic tweezers, there are some differences between the two. In our device, trapping was accomplished by utilizing outer streaming flows (the microvortices), beyond the Stokes layer (∼6 µm), see Figure 4, instead of the inner eddies. With the rotational velocity under of ∼150 µm/s (0.5 Hz at 3 Vpp), the shear stress on cells is estimated to be ∼1 × 10-2 N/m2, as modeled by a vortex flow matching the rotational velocity data (Figure 5). It increases to 4 × 10-2 N/m2 at the higher rotational speed of ∼600 µm/s (3 Hz at 7 Vpp). These values are considered small; hence, stress on cells is not likely to adversely affect the bioparticles. Figure 8 demonstrates trapping of nanoscale bioparticles (antibodies). Results are presented in similar format as that of figure 7. Figure 8a shows the trajectories of antibodies entering, being trapped, and leaving the microvortices. Since the antibodies are 3 orders of magnitude smaller than cellular objects, observation of an individual antibody is not possible and only their scattering of small fluorescent spots can be observed. Figure 8b presents the results on trapping force upon the antibodies. The maximum trapping force (gray area) increases with Reynolds number of microvortices, which is in accord with that discussed in Figure 7b. The maximum trapping force exerted by the microvortices is in the range of 160 ± 50 fN. Although, strictly speaking, Stokes law can only be applied on spherical objects, nonspherical particles can also use the Stokes law to approximate the spherical-equivalent diameter. Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

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The difference in calculated Stokes drag between a spherical body and a nonspherical one is as follows: 15% for a circular disk with its surface facing the flow; 43% for the same disk with surface parallel to flow; 40% for elliptical body of revolution (1:3 aspect ratio) with axis parallel to flow; and 80% for the same body with axis normal to flow.29 Even though it is known that the geometry of antibody is shaped like a “T” or “Y”, details of its orientation to the flow is not known and hence would contribute substantially to the uncertainty. The equivalent Stokes diameter of IgG we used is 10 nm and is perhaps the best estimate for the scenario. In an case, the inaccuracy incurred in adopting the effective Stokes diameter approach is comparable to the measurement error (see Figure 8b). Comparison of results of the trapping force between cell-sized particles (Figure 7) and that of nanosized ones (Figure 8) reveals an intriguing fact: the 10-µm cell-size particles give rise to a force level of ∼10 pN where 10-nm antibodies results in ∼100 fN force. Based on the results of the cell-size particles, one would expect the trapping force of antibodies to be 10 fN, not 100 fN. This 10fold increase in trapping force suggests that the microvortices can retain small macromolecules much stronger than of larger cellular objects. Consequently, antibodies could still be held by the microvortices at a very large flow velocity of 1680 µm/s at 7 Vpp. One possible reason for the increased trapping ability for smaller particles is that trajectories of the nanosized antibodies cover a wider range of radius of curvaturesfrom near the vortex core to the outer rimscompared to that of the cellular particles, which were observed to have trajectories of only 3-5 cells from the inner core to outer vortex rim for a microvortex with major axis of ∼100 µm. Since the Stokes numbers for antibodies (∼10-11) and cellsize particles (∼10-5) are all small, all bioparticles tested faithfully follow streamlines on which they are upon. We believe the fundamental reason that the particles are trappedsa departure from its original streamline of the background flowsis that a stagnation point, or points, exist(s) when the flow field of the microvortices is superposed to that of the background flow. There are two likely reasons for the stagnation point: (1) since the background flow is parallel to the plane of the two vorticessnot perpendicularsat the top of one vortex (the one further away from the upstream flow) the vortex flow is opposite in direction to that of the background flow; and (2) there are boundary layers on the walls of the microchannel. Once a stagnation streamline exists, a particle can depart from its original path and follows a different streamline. From above experiments, our device does seem that it can indiscriminately trap bioparticles of all sizes testedswhen we injected a mixture of multiple particles (1, 5, 10, and 15 µm) into the device they were all trapped by the vortices. However, it is important to note that the flow rate of the background flow required to “release”, or untrap, the trapped bioparticles strongly depends on particle size. That is, much larger flow rate is required to release the antibodies than the cells (compare the right ordinates of Figures 7 and 8). There are three implications: (1) the device can be used to enrich, or concentrate, bioparticles in medium with low concentration; (2) to sort heterogeneous mixture (29) White, F. M. Viscous Fluid Flow, 2nd ed.; McGraw-Hill: New York, 1991; pp 178-179.

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of bioparticles via the aforementioned effect of background flow on trapping; (3) a combination of the above. Trapping of Nonspherical Bioparticles. To scrutinize the ability of the microvortices to trap bioparticles with variation in shape, trapping of RBCs were tested with a microchannel 50 µm in height. The disk-like cells are far from spherical, with the major diameter of ∼8 µm and distance across the disk near the center of ∼1 µm. RBCs are also very deformable as they transport through small vessels. These properties, and their availability, make RBCs ideal to explore the trapping of nonspherical bioparticles in our device. Results showed that RBCs are trapped in the same manner as cell-size particles tested, see Figure 9a, with indistinguishable overall difference. However, a tumbling motion of RBCs was observed as they were entering and leaving the microvortices. This motion was not seen when they were within the microvortices due to observation difficulty. Trapping of RBCs suggests that the microvortices seem to have no geometry bias in trapping bioparticles. This fact should enable the microvortices to be a versatile platform in confining bioparticles in a small region in space for bioassay studies. Cell Viability Tests. It is important to understand the effect of the circulatory motion on the cells as they are being trapped to ensure that their physiological state is not compromised. Toward this end, a cell viability test of HEK cells was performed with calcein AM fluorescence stain. With PBS medium in the microchannel at 20 µm/s, HEK cells were injected into the device and allowed to be trapped continuously for 30 min. Afterward, calcein AM were perfused into the microchannel to stain the trapped HEKs. Fluorescence results in Figure 9b show all trapped cells were stained positively, suggesting the cells were viable with no perceptible damage. The test was performed with the device bonded with a microchannel of 50 µm in height. Capacity of a Microvortex. The capacity of a microvortex to trap microparticles depends on the type of bioparticles and the dimension of the microvortex as determined essentially by the channel height. The capacity of a microvortex to trap particles depends on the type of bioparticles and the dimension of the microvortex, which is essentially determined by the channel height. For 50 µm channel height, 1 to 5 cell-size bioparticles can be trapped within a microvortex. However, for red blood cells, several tens of cells can be trapped. For IgG antibodies, it was difficult to count but more than that of red blood cells. For 1000 µm channel height, 10-20 cell-size particles can be trapped. CONCLUSIONS This study presents a noninvasive, hydrodynamic approach to confine suspended bioparticles within a spatial region as the background flow passes and their subsequent controlled release. Microvortices generated by in-plane resonating motion (140 kHz) of a microplate are leveraged to trap the bioparticles. The rotational frequency of the microvortices can be robustly controlled and ranges from 0 to 6 Hz under driving voltage of 2-9 V (peak to peak). Successful trapping were performed with polystyrene beads (10 µm), RBCs, HEK cells, and IgG (10 nm) under a range of background flow, with a maximum of 1680 µm/s for antibodies. The trapping force of the microvorticessa measure of the ability of a microvortex to retain objects rotating along its circulatory streamlinesswere found to achieve a maximum of 12

Figure 9. Results with RBCs and HEK cells. (a) Trapped RBCs (dark ring) tracing a rotating trajectory above the edge of the oscillatory plate. (Only a few cells are trapped on the right-side microvortex; hence, the dark rotating ring is not as evident.) (b) Fluorescence micrograph of cell viability test on trapped HEK cells. (Dashed lines show the oscillatory plate.) Calcein AM was supplied into the channel after cells were trapped for 30 min. (Upper left region (i) shows three bright green fluorescent marked cells adhering to the bottom of channel.) A cluster of HEK cells in region (ii) are being trapped by the two microvortices. All cells are stained positively with calcein AM, which proved cells in vortices are viable. The device was bonded within a 50 µm height microchannel and operated at 3 Vpp. A movie is available in the Supporting Information.

± 2.0 pN for cell-size particles (10 µm) and 160 ± 50 fN for nanosize antibodies. Trapping of nonspherical RBCs was also successfully demonstrated. Cell viability tests of HEK cells after being trapped for 30 min prove cells are viable, suggesting the microvortices are harmless to the cells as a trapping tool. The volume available of a microvortex to trap particles is essentially constant with fixed channel geometry. Based on the orbit of trapped bioparticles, the trapping volume is calculated to be ∼10 nL for an average 90-µm-diameter microvortexspresent in large container (used in Figure 4) and 1000-µm-height channel (used in Figures 5, 7 and 8). This volume decreases to ∼5 nL for the 50-µm-height channel used in trapping of RBCs and HEK cells (Figure 9). In terms of cell count, a 90-µm-diameter microvortex would trap 10-20 cells. At a channel height of 50 µm, the vortex diameter was constrained to ∼40-µm average diameter, which results in trapping of maximum of only ∼3 HEK cells (diameter ∼15 µm) and ∼10 RBCs (average diameter ∼8 µm). Boundary layer effect on rotating cells is likely to have an additional effect on limiting the number of trapped particles, but is difficult to quantify. Nevertheless, the constant-volume characteristic of the microvortices is deemed beneficial since, in most applications, the size of bioparticles is known and thus the count can be estimated. We cannot precisely control how many particles are trapped; in almost all cases, multiple particles are considered, since the trapping volume allows for multiple particles. Hence, in the force measurement experiment (Figure 7) multiple particles are involved during the trap/ release process at a particular voltage and background flow. However, the particles, especially cells, might not be trapped/released at the same time due to slight variation in size. There are some potential advantages to this new hydrodynamic trapping approach. First, the shear stress in trapping cell-size bioparticles (∼10-2 N/m2) is much less than other noninvasive methods and should ensure gentle handling of biosamples. Second,

the microvortices should not be sensitive to difference in fluid mediumse.g., DI water, PBS buffers, or other biological fluidsssince the driving mechanism is medium-independent, although a viscous effect from different fluids might cause rotational velocity or amplitude variation. Third, the trapping force is higher per diameter of bioparticle for nanometer-scale bioparticles than cell-size ones. Fourth, once trapped, the concentration of bioparticles increases with time performing the task of enrichment. This function is more pronounced with smaller bioparticles, e.g., antibodies, than larger ones since microvortices have an essentially fixed trapping volume. Fifth, the microvortices do not appear to have a geometric bias in trapping, as demonstrated by trapping of RBCs. Sixth, minimal external equipment support, e.g., no laser as in optical tweezers, is needed. Future applications of the technique might include various bioassay platforms, such as cell culture of suspended cells after being trapped, drug screening via drug-containing medium flowing past trapped cells, and enrichment of low volume fraction bioparticle studies. ACKNOWLEDGMENT Funding support of this work through grants NSC 95-2120M-002-006 and NSC 96-2120-M-002-002 from the National Science Council of the Republic of China is gratefully acknowledged. The help of Prof. H. Lee, Department of Life Science, National Taiwan University, in working with bioparticles is also appreciated. SUPPORTING INFORMATION AVAILABLE Two movies corresponding to Figure 4 and 9b. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review May 11, 2008. Accepted August 27, 2008. AC800972T

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