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Trapping of Bioparticles via Microvortices in aMicrofluidic Device for Bioassay Applications

Cheng Ming Lin, Yu Shang Lai, Hsin Ping Liu, Chang Yu Chen, and Andrew M. WoAnal. Chem., 2008, 80 (23), 8937-8945 • Publication Date (Web): 29 October 2008

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Trapping of Bioparticles via Microvortices in aMicrofluidic 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 bioparticlesin a microfluidic device. An in-plane oscillatory micro-plate, driven via Lorentz law, generates two counter-rotating microvortices, trapping the bioparticles within theconfines of the microvortices. The force required to trapbioparticles is quantified by tuning the background flowand the microplate’s excitation voltage. Trapping andreleasing of 10-µm polystyrene beads, human embryonickidney (HEK) cells, red blood cells (RBCs), and IgGantibodies were demonstrated. Results show the micro-vortices 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 thebackground flow exceeds a limit. This flow limit increaseswith the rate of rotation, which defines the trap/releaseforce boundary over the range of operation. This boundaryis 12 ( 2.0 pN for cell-size bioparticles and 160 ( 50 fNfor antibodies. Trapping of RBCs demonstrated micro-vortices’ ability for nonspherical cells. Cell viability wasstudied via HEK cells that were trapped for 30 min andshown to be viable. This hydrodynamically controlledapproach to trap a wide range of bioparticles should beuseful as a microfluidic device for cellular and subcellularbioassay applications.

The lab-on-a-chip approach leverages upon microfluidics tech-nology1 and is becoming an enabling platform for miniaturizationof biological and chemical analyses. Characteristically, thesedevices require small sample volume, fabricate with relative ease,are conveniently controlled,2 and often utilize rapid detectionprocess.3 They provide a suitable environment for studies ofbiosamples while integrating various analytical operations in aperfusion system.4 For example, microfluidic system is capableof real-time bioassays of controlling fluid conditions, includingexchange of fluid media,5 supply of sufficient grow factors, andrefresh fluid for live cells and biomolecules.6,7

In cellular studies via microfluidics, some form of cell-positioning method is often needed. Invasive (contact) cell

trapping can provide direct control but faces challenges due tothe complex physical properties of biosamples; for instance, whiteblood cells are very sticky while RBCs are rather nonadhesive.Carlson et al. have designed mechanical filters for trappingdifferent cell types due to their different physical properties.8

Moreover, noncellular entities with similar properties to cells weretrapped 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 ap-proaches often cause contamination to biosamples and result inone-time usage of the device.

A noninvasive manipulation method to control cells is prefer-able. An ideal cell diagnostic platform should retain the naturalproperties of cells and, if needed, provide a highly efficientmedium exchange with limited side effects to cells. Such mediumexchange can supply nutrients and remove metabolites, along withensuring a stable and gentle environment for live cells.12 Cellmanipulating using a noninvasive approach is the best approachto 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 focusedlaser beam to generate forces on a cell based on the difference inthe refractive index between the cell and the medium. Trappingis achieved by focusing the laser beam through the microscopeobjective. Sylvie Henon et al.18 used the OT to hold a red blood

* 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.

(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.

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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.

Anal. Chem. 2008, 80, 8937–8945

10.1021/ac800972t CCC: $40.75 2008 American Chemical Society 8937Analytical Chemistry, Vol. 80, No. 23, December 1, 2008Published on Web 10/29/2008

cell (RBC) and determined the shear modulus of cellularmembrane.

Dielectrophoresis (DEP) can trap polarizable bioparticles undera nonuniform electric field. The approach has been well studieddue to its relative ease of fabricating planar electrodes in a myriadof configurations, each resulting in a unique electric field for aspecified task. Voldman et al.15 have shown that a negative DEPconfiguration induced effective dipole moment in a cell that isantiparallel to the electric field, which creates a dielectrophoreticforce that could trap the cell stably. Also, Becker et al.19 showedthat breast cancer cells could be attracted to electrode tips viapositive DEP traps, suggesting proper design of electrodes cantrap and move the cell to a predetermined position. However,polarization of cells induced by electric field may affect theirinterior. Perhaps the most severe limiting factor for utilization ofDEP is the need to employ suitable combination of medium andcell, based on their dielectric (not physiological) properties, inorder to produce a desired manipulative force.

Another noncontact method is acoustical tweezers, whichgenerates a trapping force via the difference in compressibilitybetween a cell and the medium. The induced pressure differenceaggregates the cells by controlling the acoustical energy. Thistechnique usually uses an ultrasonic standing wave to trap particlesand cells.20,21 It provides a rapid and reliable trapping quality forcells and particles.13,20 Ultrasound has been used for many yearsin the studies of microscale particles. By changing operationfrequency and reflecting distance, the trapping forces can becontrolled and specific particle sizes can be trapped.

There have been several noninvasive hydrodynamic-basedstudies to trap cells in recent years, with one of the maindifferences in the method of generating the needed trappingmechanism. Shelby et al.22 have utilized microvortices to manipu-late bioparticles and measured their rotational rate within aconfined region, and application of the technique on nanoparticleshas been demonstrated.17 The flow in which cells were trappedwas induced in a novel manner by the main channel (steady)flowsthrough an opening in the channel wallsleading to second-ary flow in the cavity where cells were located. Lutz et al.23 werethe first to utilize secondary streaming flow to trap cells behinda fixed cylinder and measured the motility force of motile cells.The streaming flow in this hydrodynamic tweezers23 approachwas generated by an external piezoelectric apparatus. Hence, onecan quantify the trapping force by varying the frequency oramplitude of the streaming flow. Although these excellent workshave provided progress on hydrodynamic trapping, much workis still needed in exploiting the full potential of the advantages ofthe method, e.g., providing a gentle environment for biosamples.

In this study, hydrodynamic trapping of bioparticles wasdemonstrated in a microfluidic device utilizing a resonatingmicroplate driven by Lorentz force, generating two counter-rotating microvortices. After quantifying the characteristics of the

microvortices, studies were conducted on the force required totrap/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 cellcondition after being trapped to ensure utilization of themicrodevice.

DESIGN OF THE DEVICEOverall Concept. The force acting on a suspended bioparticle

under some local flow condition is proportional to the fluidproperties, the bioparticles’ geometry, and the velocity differencebetween the bioparticle and the flow. If a desirable local flow canbe generated, the resultant hydrodynamic force might be suitablefor manipulating bioparticles, avoiding the undesirable effects ofphysical contact methods.

In mesoscale flow, a vortex is often generated via relativelyhigh speed flow over a sharp corner or a cavity. As such, asuspended particle could enter the vortex and be trapped withinthe 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 utilizingthem to trap bioparticles.

Design of Microvortices Generator. Lorentz force wasutilized to drive a microplate to resonance (140 kHz). Oneadvantage of using Lorentz force is to reduce the complexity ofactuation of the MEMS-based device by employing an externalmagnet. Once in resonance, the microplate generates two micro-vortices near its edges, providing a local flow condition forhydrodynamic manipulation of bioparticles.

The main structure of the vortices generator consists of adouble-clammed, suspended bridge, combined with a square platein the middle as the primary structure, as shown in the 3D sketchof Figure 2a. When an alternating current (ac) passed throughthe gold layer on the surface of the suspended bridge in thepresence of an external magnetic field B (Nd-Fe-B magnet, ∼1T) perpendicular to the bridge surface, the main structure wasforced to resonate in the third (in-plane) direction. The geometryof the bridge is 1.2 µm thick, 20 µm wide, and 750 µm long, withthe 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 drivingsource is embedded within the present device versus external.23

Second, the region where cells are trapped is in the outerstreaming flow in this study versus within the inner eddies. Third,

(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.

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

8938 Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

the frequency of oscillation was fixed at 140 kHz versus 40 Hz to1 kHz. These differences have consequences in the entire flowfield and, more importantly, on the trapped bioparticles, e.g., shearstress, as will be discussed below.

Quality Factor. This vortices generator is a resonant-basedactuator operating under liquid with the natural frequency ∼140kHz. This is implied from the rotational velocity of the vortices,which maximizes at this frequency and decays substantially atoff-resonance frequencies. The maximum displacement of theoscillatory plate is less than 1 µm. From these characteristics,the quality factor (Q) is calculated to be ∼10, which is indicativeof a reasonable resonating device.

EXPERIMENTAL ASPECTSMicrofabrication. Fabrication of the suspended structure

utilized conventional lithographic microfabrication, as shown inFigure 2b. First, silicon nitride (∼1 µm) was deposited on thesilicon substrate with a low-pressure chemical vapor depositionsystem. 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. Unnecessarymetal was removed by metal etchant. Next, the positive photoresist

on the metal is removed by acetone. By repeating the procedureof coating photoresist and development, the open window of thecavity underneath the oscillating structure was formed. The siliconnitride not covered with the photoresist would be etched usingRIE. Then, a bulk micromachine process was used to removeunwanted silicon below the plate/beam structure using potassiumhydroxide, thus suspending the structure. Only two masks areneeded in the fabrication procedure.

The microchannel was fabricated with poly(dimethylsilane)(PDMS, Sylgard 184, Dow Corning) via a soft lithographic tech-nique.24 The ratio of elastomer/curing agent used was 10:1 by weight,mixed uniformly and degassed at low pressure for 20 min. Next, thepremixed PDMS was poured on a master and baked at 85 °C for1 h. After cutting and peeling off from the master, the replica PDMSchannel is formed. There are two different PDMS channel height inthis work: (1) 1000 µm for characterizing microvortex and trappingforce (used in Figures 5, 7, and 8 and (2) 50 µm for RBCs and cellviability test (Figure 9).

Device Assembly. The integrated microfluidic device iscomposed of a silicon chip (28 × 20 × 0.5 mm3) that houses thevortices generators, a PDMS microchannel, and a poly(methylmethacrylate) (PMMA) board (50 × 50 × 3 mm3), as shown inFigure 3. A linear array of four vortex generators in a silicon chipwas mounted on PMMA for easy handling. A PDMS channel wasbonded to the surface of the silicon chip using an oxygen plasmatreatment. Polyethylene tubings (1.09-mm o.d. and 0.38-mm o.d.)were connected to fluid connection components on the back sideof PMMA, providing a convenient method for cell and mediuminjection by a syringe pump. PMMA board includes three drilledholes: two inlet holes and one outlet. A permanent magnet wasplaced directly beneath the oscillators, with it large enough thatthe magnetic field can be considered uniform for all oscillators.

Procedures. The chip and other parts of the microfluidicdevice were first sterilized using 70% ethanol. Then, bioparticlesspolystyrene beads, cells, and antibodiesswere injected into thedevice to study the trapping characteristics of the microvortices.Further details are provided in subsequent sections.

For trapping of cell-sized bioparticles (10 µm), an opticalmicroscope was used to observe the process of vortex generationand to study the trapping/releasing phenomenon. Additionally, aCCD camera (Unibrain, Fire-i400) recorded images for postpro-cessing, at the maximum frame rate of 30 frames/s at resolutionof 640 × 480. The interface IEEE1394 provides sufficient band-width for data transmission from CCD to PC.

To probe the trapping performance of bioparticles smaller thancellular level, biomolecules with the effective Stokes diameter of10 nm were studied. The trapping experiment was done via thesame device and measurement procedure but the operation wasobserved with an inverted fluorescent microscope (Olympus IX-71) with a CCD camera (Olympus DP-70).

Bioparticles and Preparation. Biological particles varygreatly in dimensions and shapes, which might affect the effective-ness of their trapping via the microvortices. Hence, a range ofbioparticles were studied to scrutinize the generality of usage ofthe device. Bioparticles tested included the following: 10-µmpolystyrene beads, human embryonic kidney (HEK-297T, ATCC)

(24) Xia, Y. N.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37 (5), 551–575.

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

8939Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

cells, RBCs, and goat anti-rabbit IgG (Alexa Fluor 546, MolecularProbes, Eugene, OR, Catalog No. A-11010). The cell-size polysty-rene beads analogically performed as cells trapped by themicrovortices for the initial observation. The HEK cells repre-sented behavior of living cells within the hydrodynamic environ-ment of the microvortices and allowed testing of cell viability. TheRBCs are nonspherical in shape and thus serve to characterizethe efficacy of the microvortices in regard to shape dependency.Finally, the use of much smaller (∼10 nm) biomolecules wouldtest the extent of generality of trapping/releasing claim of thedevice.

Details of preparation of HEK cells and RBCs are provided asfollows. HEK cells were grown in Dulbecco’s modified Eaglemedium (Gibco/Invitrogen) supplemented with 10% fetal bovineserum (Gibco/Invitrogen). Then, 1% penicillin-streptomycin and1% sodium pyruvate were added at 37 °C in a humidifiedenvironment of 5% CO2 in air. Prior to injecting into the microde-vice, cells were detached from the culture dish, treated in trypsin-ethylenedinitriletetraacetic acid (trypsin-EDTA) in a 37 °C incu-bator for 4 min, and then resuspended in medium. Cells andmedium were then centrifuged at 1000 rpm for 5 min. Thesupernatant was aspirated, and cells were resuspended in 10 mLof PBS solution with a pH of 7.4. RBCs were separated from wholeblood cells by centrifugation and, to alleviate clogging, dilutedwith PBS, achieving a density of 104 cells/µL.

Cell viability test was conducted to ensure the microvorticesplatform is indeed appropriate for cellular studies. Acetomethoxyderivate of calcein (calcein AM, C3100MP, Invitrogen) wasprepared in stock solution (50 µg of calcein AM in 9.6 µL ofDMSO) and diluted with standard phosphate-buffered salinesolution (PBS) in a volume ratio of 250. After HEK cells weretrapped for 30 min, calcein AM was perfused into the channel,testing the cells for viability while cells were still rotating withinthe microvortices. Calcein AM can transport through the cellularmembrane into cells then be hydrolyzed by an intracellularenzyme resulting in strong green fluorescence. As dead cells lackthis enzyme, only live cells are marked. Fluorescent images weretaken via the CCD camera (DP-70, Olympus) using an invertedmicroscope (IX-71, Olympus).

Temperature Measurements. Generation of the microvor-tices requires an electric current flowing through the device,which results in fluid temperature increase. This temperatureincrease in microchannel can be considered theoretically as powerdissipation from the oscillator surface when a current passingthrough a material, i.e., P ) V2/R, where I and V are the root-mean-square values of current and ac voltage, respectively.

Exposure of cells to high temperature would cause extra stressand results in irreversible damage. To probe the effect of tem-perature increase with various driving voltages, the fluid temper-ature within the microchannel was quantified at two typicalbackground flow velocity, i.e., 20 and 100 µm/s.

Characterization of temperature effect was conducted asfollows. Temperature-dependent fluorescent dye, Rhodamine B(Sigma-Aldrich, Inc.), was used since its response is sufficientlyrapid (ms) and with good spatial resolution (µm).25 With thistechnique, fluid temperature can be evaluated via fluorescentintensity, which would decrease as temperature increases. Asolution of 0.1 mM Rhodamine B was injected into the microdeviceand imagine acquisition was performed by a CCD camera (DP-70, Olympus) mounted on an inverted fluorescent microscope (IX71 Olympus, Hg lamp, 10× objectives). To calibrate the intensitywith temperature, a thermal couple was placed in contact withthe back side of the silicon chip bonded with PDMS channel. Aheater was embedded in a Petri dish in which our device wasplaced upside down lying on the bottom of the dish. Then, thedish was filled with water in order to provide a stable temperaturereservoir as the heater was activated. Fluorescent imagine of stabletemperature from 25 to 70 °C was recorded. The calibration curvewas generated by averaging the gray value of intensity corre-sponding to each temperature. By infusing the fluorescent dyeinto the activated microdevice, we can evaluate the temperaturein the vicinity of the microvortices. Four sets of data were takenat each driving voltage. The gray value at room temperature wassubtracted from the mean gray value of the intensity data at eachtemperature in order to reduce the environmental variations overtime, such as lamp intensity. The standard variation of thetemperature increase was less than 1 °C. The experiments wereconducted for both 20 and 100 µm/s background flow velocity.

Hydrodynamic Force. In this work, bioparticles were hydro-dynamically trapped via the microvortices. Procedurally (seeFigure 7a), bioparticles were pumped into the device upstreamof the vicinity of the microvortices, with them rotating relativelyrapidly. This ensures the bioparticles are trapped by the micro-vortices and rotate with the vortices while the background fluidpasses by. Then, the background flow velocity was reduced, alongwith decrease in the excitation voltage until a particular voltageto be tested was reached. With the bioparticles still trapped androtating with the microvortices, the flow was gradually increaseduntil they were released. The entire process was recorded by CCDcamera for postprocessing. The procedure was repeated for other

(25) Ross, D.; Gaitan, M.; Locascio, L. E. Anal. Chem. 2001, 73 (17), 4117–4123.

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

8940 Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

excitation voltages. The device operated at a frequency of 140 kHzand voltage of 2-7 Vpp.

The trapping force of the microvorticessa measure of theability of a microvortex to confine a bioparticle rotating along its

circulatory streamlinesswas quantified. If the hydrodynamicforce from the background flow acting on a bioparticle exceedsthe maximum trapping force corresponding to a particular vortexrotational velocity, the bioparticle would be released from themicrovortex, and conversely, a force from the background flowless than the maximum trapping force would ensure that thebioparticle remains in the trapped state. Thus, this maximumtrapping force can calculated from the classical Stoke’s law foran object immersed in low Reynolds number flow, i.e.,

Fmax ) 6πηUa (1)

where η is the viscosity, U is the velocity of the background flowthat would cause release of a trapped bioparticle, and a is theradius of the bioparticle.

RESULTS AND DISCUSSIONMicrovortices. The primary result of the in-plane resonating

motion of the microplate is generation of two counter-rotatingmicrovortices. Figure 4a presents a micrograph of the microvor-tices, showing time-lapsed trajectory of microvortices, made vis-ible by trapped 10-µm polystyrene beads. Figure 4b sketchesthe trajectories based on postprocessing of dynamic image data.The trajectory indicates the flow within the vortex approaches theoscillatory plate from above, moves outward along the plate for ashort distance, then leaves the plate area and curves upward, andeventually progresses downward toward the plate, completing therecirculating flow circuit. Once trapped, particles recirculatecontinuously within the confine of the microvortices just abovethe 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 outerrim. The width of orbit appears to depend on the uniformity ofthe trapped particles: 1-2 diameters for 10-µm polystyrene beadsand 2-3 cell diameters for, say, HEK cells. However, the numberof bioparticles entering each microvortex might not be the same,which is the case in Figure 4a. The microvortices are reliable androbust. The diameter of the microvortices ranges from 80 to100 µm.

Figure 4. Counter-rotating microvortices. (a) Micrographs, takenfrom snapshots of movie frames (using a portable digital microscope,with a frame rate of 15 frames/s), showing time-lapsed trajectoriesof 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 ofthe 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 ofthe oscillatory plate, as observed from Figure 4a. The orbits of themicrovortices 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 rightordinate. Results indicated the rotational velocity increases paraboli-cally with voltage. The microvortex is very controllable and robust.Statistic analysis performed using Student’s t tests at 95% confidencelevel, n ) 3, S ) 1.0. The regression coefficient is 0.97.

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

8941Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

Effect of Particle Size. Intuitively, particle size should affectthe efficacy of the microvortices in confining the particles withinthe region of the recirculating flow since trapped bioparticles needto faithfully follow the elliptical streamlines of the microvorticesagainst the possibility of being washed downstream by the flowingmedium. Thus, a smaller particle should be more readily trappedthan a larger one. To test this hypothesis, different sizes ofpolystyrene beadss1, 5, 10, and 15 µm in diameterswere testedin the microfluidic device. Trapping was commenced whensuspended particles of a particular size flowed near the trappingzone and into the microvortices above the oscillatory plate. Resultsshow 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 ofsmaller particles. The 1-µm particles are sufficiently small thatare very well confined within the microvortices. This sizeindependency characteristic suggests that the microvortices mighthave general application as a trapping tool.

Rotational Velocity of Microvortices. Rotational velocity ofthe microvortices should play a key role in defining the overalltrapping characteristics since trapped particles are confined withinthe region of the microvortices, advecting along in elliptical orbit.Dimensionally, one expects rotational velocity depends on theoscillatory frequency of the microplate and a length scalesmicroplate dimension, viscous penetration depth, or amplitude ofoscillation. The data will provide clues to which length scale iscorrect.

Figure 5 presents the results of rotational velocity for a rangeof driving voltage of the microplate. The average rotational velocityis in the order of hundreds to 1000 µm/s, which is a relativelyfast value in microflow environment, and varies quadratically withdriving voltage, having a threshold voltage of 2 Vpp. The secondordinate shows the rotational frequency of the microvortices, withmaximum frequency of ∼6 Hz at 9 Vpp. Much smaller antibodies(10 nm) were also used as tracer particles in the rotational velocityexperiment. Results showed the relationship between rotationalvelocity and driving voltage to be essentially equivalent to that ofusing polystyrene beads. In this work, with the geometry of themicroplate and the frequency fixed, the variation of rotationalvelocity with voltage is largely due to variation of oscillatoryamplitude with voltage (also see Schlichting28).

It is interesting to note that the theoretical magnitude of therotational (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, ∞))-34

U0

ndU0

dx(2)

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

increase in Lorentz force causes an increase in the plate’samplitude, and (2) the increase in temperature lowers theviscosity, which would further increase the amplitude due todecrease in viscous damping. Hence, both effects would contributeto the rotational velocity versus voltage data.

Temperature Increases. The measured temperature increaseby Joule heating showed the quadratic relation to drive voltageas we expected (see Figure 6.). Here, temperature rise is definedas the increase from 0 V condition, which is essentially at roomtemperature (25 °C). Two regression models of second orderexpress the temperature response of fluid relative to roomtemperature under two different flow velocities in a microchannel.At drive voltage of 3 Vpp, the temperature increase is under 5 °Cand the trapping force is good at 10-µm particles when the suppliedbackground flow velocity was at 20 µm/s, producing an absolutetemperature of 30 °C in the microchannel when room temperaturewas at 25 °C. With increasing background flow velocity to 100µm/s, the temperature rise was only 2 °C. This temperatureincrease 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., trappingdata (in Figure 7) correspond to 2-7 Vpp only. This is becausethe temperature within the microdevice at 8V is beyond physi-ological temperature of 37 °C. However, measurement of rotationalvelocity was performed up to 9 Vpp.

Trapping Force. Figure 7 presents the trapping force of themicrovortices on cellular-sized particlessrepresented by measure-ment of 10-µm polystyrene beadssover a range of parameters.The trapping force is determined at the point that a vortex at a

Figure 7. Trapping force measurement of cellular-scale micropar-ticles. (a) Micrograph (top view) showing trajectories of particlesentering, being trapped (black spots), and leaving the microvorticeswhen the hydrodynamic force exerted by the background flow on thetrapped particles is greater than the trapping force of the microvor-tices. (b) Trapping force of microvortices on 10-µm polystyrene beadsunder a range of parameters. The x-axis is the Reynolds number ofthe counter-rotating microvortices, expressed as ReΓ ) Γ/η, where Γ) IV ·dl, the circulation of a vortex. The operating voltage for trappingforce measuring is from 2 to 7 Vpp. The Reynolds number scale (Ref

) U ·h/η) on the far right ordinate characterizes the flow in themicrochannel. The gray region represents the boundary betweentrapped and untrapped regions. Mean value and error bar range werecalculated using Student’s t tests with 95% confidence level.

8942 Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

particular rotational velocity can no longer retain a trapped particleagainst a particular background velocity, thus releasing it down-stream. As shown in Figure 7a, this trapping force was quantifiedvia a three-step process: first, observing particles that are embark-ing upon the trajectory to enter the microvortices; second, trappingparticles at a particular vortex rotational velocity, i.e., at a particulardriving voltage; third, releasing trapped particles by increasingthe background flow velocity, while keeping the voltage fixed,until the particles leave the microvortices and progress down-stream along the microchannel. This process was done with singleor multiple particles (1–3, but more can be trapped). The point ofrelease usually happens at the top of the microvortex far fromthe plate. By recording images of this process by a CCD camera(30 frames/s), the background flow velocity at which releaseoccurs is fairly easy to determine.

Figure 7b presents the results of the trapping force. Theabscissa represents the Reynolds number of the microvortices,expressed as ReΓ ) Γ/η, where η is the kinematic viscosity andΓ ) I Vb ·dLb is the circulation of the vortex at mean rotationalvelocity (V) (from Figure 5) along the recirculation path (L), or∼300 µm for 100-µm-diameter microvortices. Thus, the circulationcorresponds directly with the driving voltage (see Figure 5). Thefar right ordinate characterizes the flow in the microchannel,represented by the Reynolds number Ref = U · h/η, where U isthe background flow velocity and h the microchannel height.Results show a distinct trend of an increase in trapping force withincreasing Reynolds number of microvortices, with the force levelin pico-Newton range. The gray area corresponds to the maximumtrapping force, or Fmax in eq 1, and divides the results into tworegions, trapped or untrapped, and can be understood as follows.At a particular Reynolds number of microvortices, a particle iswell trapped under low Reynolds number of background flowcondition (below the gray area); hence the trapping force issufficient to retain the particle within the trapped region. As thebackground flow increases (above the gray area), the particleescapes the microvortices, entering the untrapped region. Alter-natively, at a particular background flow, a particle is not trappedat low microvortex rotational velocity (left of the gray area) andbecomes trapped when the microvortex velocity is increased (rightof the gray area). Thus, at a specified Reynolds number ofbackground flow (Ref), particles can be trapped at a range ofReynolds number of microvortices (ReΓ) above the value definedby the gray area corresponding to that value of Ref. The maximumtrapping force measured was found to be 12 ± 2.0 pN at the flowvelocity of 140 µm/s and driving voltage of 7 V (peak-to-peak).

It should be instructive to compare this force level with othernoncontact methods. The cell-trapping force generated by opticaltweezers is ∼100 pN,14,26 dielectrophoretic force in the range of200-500 pN27 and acoustics 300-600 pN.13 The hydrodynamictweezers23 can provide trapping force up to 30 pN, and the shearstress on cells is stated to be comparable to arterial shear stressin circulatory system, which is ∼1.5 N/m2. These above forcesare that reported in the cited literature and should be capable of

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 onhydrodynamic tweezers23 in that the use of microvortices herefor trapping is a form of hydrodynamic tweezers, there are somedifferences between the two. In our device, trapping was ac-complished by utilizing outer streaming flows (the microvortices),beyond the Stokes layer (∼6 µm), see Figure 4, instead of theinner 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 rotationalvelocity data (Figure 5). It increases to 4 × 10-2 N/m2 at thehigher rotational speed of ∼600 µm/s (3 Hz at 7 Vpp). Thesevalues are considered small; hence, stress on cells is not likely toadversely affect the bioparticles.

Figure 8 demonstrates trapping of nanoscale bioparticles(antibodies). Results are presented in similar format as that offigure 7. Figure 8a shows the trajectories of antibodies entering,being trapped, and leaving the microvortices. Since the antibodiesare 3 orders of magnitude smaller than cellular objects, observa-tion of an individual antibody is not possible and only theirscattering of small fluorescent spots can be observed. Figure 8bpresents the results on trapping force upon the antibodies. Themaximum trapping force (gray area) increases with Reynoldsnumber of microvortices, which is in accord with that discussedin Figure 7b. The maximum trapping force exerted by themicrovortices is in the range of 160 ± 50 fN.

Although, strictly speaking, Stokes law can only be appliedon spherical objects, nonspherical particles can also use theStokes law to approximate the spherical-equivalent diameter.

(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 themicrovortices (left dotted line) and upon release (right dotted line)when the flow of the PBS medium is excessive. Enrichment ofantibody is demonstrated via increased fluorescence intensity as moreantibodies are being trapped. Black dashed lines show the suspendedplate. (b) The approximate trapping force exerting on the antibodiesby the microvortices. Axes are similar to that of Figure 7. Resultsshow the trapping force is in femto-Newton range. The operatingvoltage for measuring was from 2 to 7 Vpp.

8943Analytical Chemistry, Vol. 80, No. 23, December 1, 2008

The difference in calculated Stokes drag between a sphericalbody and a nonspherical one is as follows: 15% for a circulardisk with its surface facing the flow; 43% for the same diskwith surface parallel to flow; 40% for elliptical body of revolution(1:3 aspect ratio) with axis parallel to flow; and 80% for thesame body with axis normal to flow.29 Even though it is knownthat the geometry of antibody is shaped like a “T” or “Y”, detailsof its orientation to the flow is not known and hence wouldcontribute substantially to the uncertainty. The equivalentStokes diameter of IgG we used is 10 nm and is perhaps thebest estimate for the scenario. In an case, the inaccuracyincurred in adopting the effective Stokes diameter approach iscomparable to the measurement error (see Figure 8b).

Comparison of results of the trapping force between cell-sizedparticles (Figure 7) and that of nanosized ones (Figure 8) revealsan intriguing fact: the 10-µm cell-size particles give rise to a forcelevel of ∼10 pN where 10-nm antibodies results in ∼100 fN force.Based on the results of the cell-size particles, one would expectthe trapping force of antibodies to be 10 fN, not 100 fN. This 10-fold increase in trapping force suggests that the microvorticescan retain small macromolecules much stronger than of largercellular objects. Consequently, antibodies could still be held bythe microvortices at a very large flow velocity of 1680 µm/s at 7Vpp. One possible reason for the increased trapping ability forsmaller particles is that trajectories of the nanosized antibodiescover a wider range of radius of curvaturesfrom near the vortexcore to the outer rimscompared to that of the cellular particles,which were observed to have trajectories of only 3-5 cells fromthe inner core to outer vortex rim for a microvortex with majoraxis of ∼100 µm.

Since the Stokes numbers for antibodies (∼10-11) and cell-size particles (∼10-5) are all small, all bioparticles tested faithfullyfollow streamlines on which they are upon. We believe thefundamental reason that the particles are trappedsa departurefrom its original streamline of the background flowsis that astagnation point, or points, exist(s) when the flow field of themicrovortices is superposed to that of the background flow. Thereare two likely reasons for the stagnation point: (1) since thebackground flow is parallel to the plane of the two vorticessnotperpendicularsat the top of one vortex (the one further away fromthe upstream flow) the vortex flow is opposite in direction to thatof the background flow; and (2) there are boundary layers on thewalls of the microchannel. Once a stagnation streamline exists, aparticle can depart from its original path and follows a differentstreamline.

From above experiments, our device does seem that it canindiscriminately trap bioparticles of all sizes testedswhen weinjected a mixture of multiple particles (1, 5, 10, and 15 µm) intothe device they were all trapped by the vortices. However, it isimportant to note that the flow rate of the background flowrequired to “release”, or untrap, the trapped bioparticles stronglydepends on particle size. That is, much larger flow rate is requiredto release the antibodies than the cells (compare the rightordinates of Figures 7 and 8). There are three implications: (1)the device can be used to enrich, or concentrate, bioparticles inmedium with low concentration; (2) to sort heterogeneous mixture

of bioparticles via the aforementioned effect of background flowon trapping; (3) a combination of the above.

Trapping of Nonspherical Bioparticles. To scrutinize theability of the microvortices to trap bioparticles with variation inshape, trapping of RBCs were tested with a microchannel 50 µmin height. The disk-like cells are far from spherical, with the majordiameter of ∼8 µm and distance across the disk near the centerof ∼1 µm. RBCs are also very deformable as they transportthrough small vessels. These properties, and their availability,make RBCs ideal to explore the trapping of nonspherical biopar-ticles in our device.

Results showed that RBCs are trapped in the same manner ascell-size particles tested, see Figure 9a, with indistinguishableoverall difference. However, a tumbling motion of RBCs wasobserved as they were entering and leaving the microvortices.This motion was not seen when they were within the microvorticesdue to observation difficulty.

Trapping of RBCs suggests that the microvortices seem to haveno geometry bias in trapping bioparticles. This fact should enablethe microvortices to be a versatile platform in confining biopar-ticles in a small region in space for bioassay studies.

Cell Viability Tests. It is important to understand the effectof the circulatory motion on the cells as they are being trappedto ensure that their physiological state is not compromised.Toward this end, a cell viability test of HEK cells was performedwith calcein AM fluorescence stain. With PBS medium in themicrochannel at 20 µm/s, HEK cells were injected into the deviceand allowed to be trapped continuously for 30 min. Afterward,calcein AM were perfused into the microchannel to stain thetrapped HEKs. Fluorescence results in Figure 9b show all trappedcells were stained positively, suggesting the cells were viable withno perceptible damage. The test was performed with the devicebonded with a microchannel of 50 µm in height.

Capacity of a Microvortex. The capacity of a microvortex totrap microparticles depends on the type of bioparticles and thedimension of the microvortex as determined essentially by thechannel height. The capacity of a microvortex to trap particlesdepends on the type of bioparticles and the dimension of themicrovortex, which is essentially determined by the channelheight. For 50 µm channel height, 1 to 5 cell-size bioparticles canbe trapped within a microvortex. However, for red blood cells,several tens of cells can be trapped. For IgG antibodies, it wasdifficult to count but more than that of red blood cells. For 1000µm channel height, 10-20 cell-size particles can be trapped.

CONCLUSIONSThis study presents a noninvasive, hydrodynamic approach to

confine suspended bioparticles within a spatial region as thebackground 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. Therotational frequency of the microvortices can be robustly con-trolled and ranges from 0 to 6 Hz under driving voltage of 2-9 V(peak to peak). Successful trapping were performed with poly-styrene beads (10 µm), RBCs, HEK cells, and IgG (10 nm) undera range of background flow, with a maximum of 1680 µm/s forantibodies. The trapping force of the microvorticessa measureof the ability of a microvortex to retain objects rotating along itscirculatory streamlinesswere found to achieve a maximum of 12

(29) White, F. M. Viscous Fluid Flow, 2nd ed.; McGraw-Hill: New York, 1991;pp 178-179.

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± 2.0 pN for cell-size particles (10 µm) and 160 ± 50 fN for nanosizeantibodies. Trapping of nonspherical RBCs was also successfullydemonstrated. Cell viability tests of HEK cells after being trappedfor 30 min prove cells are viable, suggesting the microvorticesare harmless to the cells as a trapping tool.

The volume available of a microvortex to trap particles isessentially constant with fixed channel geometry. Based on theorbit of trapped bioparticles, the trapping volume is calculated tobe ∼10 nL for an average 90-µm-diameter microvortexspresentin large container (used in Figure 4) and 1000-µm-height channel(used in Figures 5, 7 and 8). This volume decreases to ∼5 nL forthe 50-µm-height channel used in trapping of RBCs and HEK cells(Figure 9). In terms of cell count, a 90-µm-diameter microvortexwould trap 10-20 cells. At a channel height of 50 µm, the vortexdiameter was constrained to ∼40-µm average diameter, whichresults in trapping of maximum of only ∼3 HEK cells (diameter∼15 µm) and ∼10 RBCs (average diameter ∼8 µm). Boundarylayer effect on rotating cells is likely to have an additional effecton limiting the number of trapped particles, but is difficult toquantify. Nevertheless, the constant-volume characteristic of themicrovortices is deemed beneficial since, in most applications, thesize of bioparticles is known and thus the count can be estimated.

We cannot precisely control how many particles are trapped; inalmost all cases, multiple particles are considered, since the trappingvolume allows for multiple particles. Hence, in the force measurementexperiment (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 thesame time due to slight variation in size.

There are some potential advantages to this new hydrodynamictrapping approach. First, the shear stress in trapping cell-sizebioparticles (∼10-2 N/m2) is much less than other noninvasivemethods and should ensure gentle handling of biosamples. Second,

the microvortices should not be sensitive to difference in fluidmediumse.g., DI water, PBS buffers, or other biological fluidsssincethe driving mechanism is medium-independent, although a viscouseffect from different fluids might cause rotational velocity or ampli-tude variation. Third, the trapping force is higher per diameter ofbioparticle for nanometer-scale bioparticles than cell-size ones.Fourth, once trapped, the concentration of bioparticles increases withtime performing the task of enrichment. This function is morepronounced with smaller bioparticles, e.g., antibodies, than largerones since microvortices have an essentially fixed trapping volume.Fifth, the microvortices do not appear to have a geometric bias intrapping, as demonstrated by trapping of RBCs. Sixth, minimalexternal equipment support, e.g., no laser as in optical tweezers, isneeded. Future applications of the technique might include variousbioassay platforms, such as cell culture of suspended cells after beingtrapped, drug screening via drug-containing medium flowing pasttrapped cells, and enrichment of low volume fraction bioparticlestudies.

ACKNOWLEDGMENTFunding support of this work through grants NSC 95-2120-

M-002-006 and NSC 96-2120-M-002-002 from the NationalScience Council of the Republic of China is gratefully acknowl-edged. The help of Prof. H. Lee, Department of Life Science,National Taiwan University, in working with bioparticles is alsoappreciated.

SUPPORTING INFORMATION AVAILABLETwo 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

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 cellviability test on trapped HEK cells. (Dashed lines show the oscillatory plate.) Calcein AM was supplied into the channel after cells were trappedfor 30 min. (Upper left region (i) shows three bright green fluorescent marked cells adhering to the bottom of channel.) A cluster of HEK cellsin 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.

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