Article pubs.acs.org/ac
Tunable “Nano-Shearing”: A Physical Mechanism to Displace Nonspecific Cell Adhesion During Rare Cell Detection Ramanathan Vaidyanathan, Muhammad J. A. Shiddiky,* Sakandar Rauf, Eloïse Dray, Zhikai Tay, and Matt Trau* Australian Institute for Bioengineering and Nanotechnology (AIBN), The University of Queensland, Corner College and Cooper Roads (Bldg 75), Brisbane, Queensland 4072, Australia S Supporting Information *
ABSTRACT: We report a tunable alternating current electrohydrodynamic (ac-EHD) force which drives lateral fluid motion within a few nanometers of an electrode surface. Because the magnitude of this fluid shear force can be tuned externally (e.g., via the application of an ac electric field), it provides a new capability to physically displace weakly (nonspecifically) bound cellular analytes. To demonstrate the utility of the tunable nanoshearing phenomenon, we present data on purpose-built microfluidic devices that employ acEHD force to remove nonspecific adsorption of molecular and cellular species. Here, we show that an ac-EHD device containing asymmetric planar and microtip electrode pairs resulted in a 4-fold reduction in nonspecific adsorption of blood cells and also captured breast cancer cells in blood, with high efficiency (approximately 87%) and specificity. We therefore feel that this new capability of externally tuning and manipulating fluid flow could have wide applications as an innovative approach to enhance the specific capture of rare cells such as cancer cells in blood.
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interactions (e.g., cells bound nonspecifically onto a surface). The ability to externally tune the magnitude of fluid shear forces (e.g., via the application of an ac electric field) provides a new capability to physically displace weakly (nonspecifically) bound cellular and molecular analytes. To demonstrate the utility of this tunable nanoshearing phenomenon, we present data on purpose-built microfluidic devices that employ ac-EHD to capture low cell numbers spiked in blood. This tunable control of surface shear forces and concomitant fluid micromixing facilitates two critical improvements to the traditional immunocapture of cellular targets: (i) enhanced capture efficiency due to an increased number of sensor−target collisions, which is a result of improved transport, and (ii) enhanced specificity resulting from the ability to tune nanoscopic fluid shear forces at the electrode interface, which serves to shear away loosely bound nonspecific species present in biological samples.
etection of specific cancer cells of interest in blood is extremely challenging owing to the nonspecific adsorption of cells and molecules present in blood samples, as many of these noncancerous cells and molecules can nonspecifically bind to the solid electrodes.1,2 While the removal of nonspecific adsorption of cells and molecules is challenging in itself, the target cancer cells are often present at concentrations millionfold lower than that of the nontarget cells (e.g., as low as 1−100 cells in 1 000 000 blood cells).3 Herein, we report a new method to avoid nonspecific adsorption of weakly bound cells and molecules using a tunable alternating current electrohydrodynamic (ac-EHD) force, referred to as nanoshearing, which drives lateral fluid motion within a few nanometers of an electrode surface. Electrohydrodynamics (EHD) deals with the fluid motion induced by electric fields.4 The use of ac-EHD to manipulate colloidal objects on the surface of flat electrodes was originally demonstrated and modeled by Trau et al.,5,6 where fluid flow arises from electrical body forces acting on f ree charges generated in solution. By manipulating the microscopic fluid flow, controlled micro- and nanostructures can be assembled, coagulated, or sheared away. This fluid flow can be further tuned by using asymmetric electrode arrays, as exploited by Brown et al.7 and others.8,9 However, to our knowledge, no one has yet attempted to use this tunable flow to preferentially select strongly bound analytes (e.g., cancer cells) over more weakly bound species which adhere to surfaces via nonspecific © 2014 American Chemical Society
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EXPERIMENTAL SECTION Reagents. Unless stated otherwise, general-use reagents were purchased from Sigma Aldrich (Australia) and immunoassay reagents were obtained from R&D/Life Technologies (Burlington, ON), Thermo-Fisher Scientific (Australia), and Received: October 4, 2013 Accepted: January 22, 2014 Published: January 22, 2014 2042
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Figure 1. (a) ac electric field E is applied to the electrode pairs, resulting in induced charges in the double layer. The characteristic thickness of the double layer is given by the Debye length,12 λD (= 1/κ), which is approximated to be 2 nm in our case (calculated on the basis of the Debye−Hückel approximation and the use of the equation given in ref 12). Upon application of E, the induced charges in the double layer of each electrode experience a force F (F = ρEt, where ρ = charge density) due to interaction with the tangential component of the applied electric field (Et) and produce a fluid flow in the direction of the broken symmetry.7,8 The asymmetric geometry of the electrodes give rise to a lateral variation in the total amount of free (double layer) charges and spatial distribution of charges on the electrode surface. Consequently, the free charges on the larger electrode create stronger lateral forces than those on the smaller electrode (FL > Fs, where FL and Fs are the resultant forces on larger and smaller electrodes, respectively), resulting in a lateral flow toward the larger electrode. (b) Reversing the polarity of the ac field also reverses the sign of the charges in the induced double layer, and since electrical body forces are the product of the charges and the applied field, a steady flow can be maintained toward the large electrode.7
maintained in Dulbeccos Modified Eagles Medium (DMEM; Gibco, UK) supplemented with 10% fetal bovine serum (FBS) and 1% pencillin/streptomycin and grown in 5% CO2 at 37 °C. MCF7 and T-47D were supplemented with 100 μg mL−1 insulin and 100 μg mL−1 sodium pyruvate. The cultured cells were trypsinized and counted by using a particle counter (Countess, Invitrogen) to obtain the desired cell density upon dilution. MCF7, T-47D, or BT-474 cells (100 000 cells/ sample) were labeled with 5 μL of DiD+ fluorescent dye (Invitrogen, UK) and incubated at 37 °C for 20 min. The desired concentration of cells was then prepared by serial dilution in PBS (10 mM, pH 7.0). Cell Capture and Detection. Whole rat blood samples were obtained from the animal house at the AIBN (UQ) and lysed as previously described.10 Briefly, cell lysis was carried out with whole blood samples centrifuged at 1000 rpm for 1 min upon the addition of RBC lysis buffer (155 mM (8.3 g L−1) ammonium chloride in 0.01 M Tris−HCl buffer, with pH 7.5). Cells were rinsed in PBS followed by centrifugation (1000 rpm for 10 min) and finally resuspended in cold PBS. Cell viability post lysis was determined using the Trypan blue exclusion assay, and approximately 97% of cells (e.g., leukocytes) was found to be viable. Subsequently, designated concentrations of DiD+ labeled target cells were then spiked into PBS, lysed blood, or 10× diluted blood (1:10 dilution in PBS). One mL of the sample was then placed into the inlet reservoirs of the devices and driven through the channel by applying ac-EHD field. The field strength was applied for 30 min with 15 min intervals (without ac-EHD) for a total pumping time of 2 h. Control experiments were performed in the absence of ac-EHD field under static conditions (without fluid flow) by filling the device with spiked samples and incubated for 2 h. Control experiments for blood samples were performed in the absence of ac-EHD field under pressure driven flow conditions using a syringe pump (PHD 2000, Harvard apparatus). To enumerate the captured cells, glass coverslips were detached from the devices and captured cells were fixed by filling the device from the top with cold methanol for 10 min, permeabilized with 0.2% Triton X-100 in PBS for 10 min, to induce cellular permeability and allow for intracellular staining.
Invitrogen (Australia). All reagents and washing solutions used in the experiments were prepared using phosphate buffer saline (PBS, 10 mM, pH 7.4). Stock solutions of antibody were diluted in PBS; photoresists for fabrication (MicroChem Corp., MA) were used as per the manufacturer’s instructions. ac-EHD Induced Fluid Flow Visualization. The small and large electrodes within the long channel of the ac-EHD micro devices (EHD-μD; Figure 1 and Figure S1, Supporting Information) were connected to a signal generator (Agilent 33220A Function Generator, Agilent Technologies, Inc., CA) via gold connecting pads. Fluid flow visualization studies were performed using fluorescent latex particles (Coulter Latron, USA) diluted in PBS (10 mM, pH 7.4). The inlet reservoirs were filled with the particle solutions, and ac field was applied using a signal generator. Fluid flow was monitored using a high speed video camera fitted onto an upright microscope (Nikon Ni-U, Japan). Device Functionalization. Prior to functionalization, the electrodes were cleaned by sonication in acetone for 5 min, rinsed with isopropyl alcohol and water for another 2 min, and dried with the flow of nitrogen. The array of gold microelectrode pairs within the capture domain of the channel were then modified with anti-HER2 using avidin−biotin chemistry (Figure 2) in a three step process. Initially, devices were incubated in biotinylated BSA (200 μg mL−1 in PBS, Invitrogen) solution for 2 h followed by coupling with streptavidin (100 μg mL−1 in PBS, Invitrogen) for 1 h at 37 °C. Streptavidin conjugated channels were then coated with biotnylated anti-HER2 (10 μg mL−1 in PBS, R&D systems) for another 2 h. The channel was flushed three times with PBS (10 mM, pH 7.0) to remove any unbound molecules after each step. Each of the surface modification steps (e.g., biotinylated BSA, streptavidin, and biotinylated anti-HER2) were performed manually by filling the microchannel with the corresponding solution to specifically modify the array of gold electrodes within the capture domain. A glass coverslip was used to seal the device after antibody functionalization. Cell Culture and Labeling. HER2(+) (MCF7, T-47D, and BT-474) and HER2(-) (MDA-MB-231) breast cancer cell lines were kind gifts of Prof. Melissa Brown (UQ), and the cells were 2043
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Figure 2. Schematic representation of cell capture and detection using an ac-EHD device.
To identify any bound WBCs, post cell capture, the devices were incubated by filling the microchannel with anti-CD45 solution for 30 min, followed by a PBS wash performed manually to remove excess anti-CD45 and subsequently stained with 4′,6-diamidino-2-phenylindole (DAPI) solution for 20 min. Images were captured using a multichannel fluorescence microscope (Nikon Ti-U upright microscope, Melville, NY) using triple stains (DiD-red, CD45-green, and DAPI-blue) and analyzed using the image processing software (Nikon Ni−S elements, Basic Research). Captured cells were counted under bright-field microscopy (Nikon Ti-U upright microscope, Melville, NY). For experiments using blood samples, nonspecific cells were recovered from the devices using TrypLE treatment at 37 °C for 5 min, and cell counts were obtained using a hemocytometer. Cell Viability-Trypan Blue Exclusion. BT-474 cells (0.5 × 105 cells mL−1) were spiked in PBS and captured under acEHD field using anti-HER2 functionalized EHD-μD devices. The captured cells were recovered from the devices using TrypLE Select (Invitrogen, USA) treatment11 at 37 °C for 5 min. 100 μL of the recovered cell suspension was transferred to an eppendorf tube and stained with equal volume of Trypan blue. Trypan blue was used to stain any dead cells, and cell counts from the stained cell suspension were obtained using a hemocytometer. Cells looking faint or dark blue within the grid were considered to be dead cells. Cell viability was obtained from the percentage ratio of the number of live cells to the total number of cells. Images of stained cells were obtained using
phase contrast microscopy (Nikon Ti-U upright microscope, Melville, NY).
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RESULTS AND DISCUSSION To investigate the nanoshearing phenomenon, we constructed two microfluidic devices (EHD-μD); EHD-μD1 contains an array of asymmetric electrode pairs within a long serpentine channel, and EHD-μD2 embraces electrodes pairs with a combination of planar and microtip spikes (see Materials and Methods and Figures S1−S3, Supporting Information). These designs facilitate large sample volume (>1 mL within 2 h) analysis that is ideally suited for clinical settings. When an aqueous solution is placed in these devices, a left-to-right unidirectional fluid flow is engendered upon the application of an ac field. Figure 1 illustrates the mechanism of ac-EHD induced fluid flow. Large and small electrodes in each asymmetric pair of the microelectrode array together form the cathode and anode (or vice versa) of an electrolytic cell. Upon the application of an ac field E, the asymmetric geometry of the electrodes gives rise to a lateral variation in the total amount of free (double layer) charges and spatial distribution of charges on the electrode surface. Consequently, the free charges on the larger electrode create stronger lateral forces than those on the smaller electrode, resulting in a lateral flow toward the larger electrode (Figure 1). A special feature of this flow is that, because all of the free charges in solution occur only within the double layer of the electrode, all of the ac-EHD body forces on the fluid also occur strictly within this region. Critically, due to the solution 2044
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Figure 3. (a,b) Capture efficiency from PBS (10 mM, pH 7.4) spiked with MCF7 (200 cells mL−1) in (a) EHD-μD1 and (b) EHD-μD2 as a function of applied frequency at constant amplitude of Vpp = 100 mV. (c, d) Capture efficiency from PBS (10 mM, pH 7.4) spiked with (c) MCF7 (200 cells mL−1 in EHD-μD1 and EHD-μD2) and (d) T-47D (200 cells mL−1) and MCF7 + MDA-MB-231 (200 cells mL−1 + 105 cells mL−1) in EHD-μD2 under ac-EHD (dark gray) and control (light gray; without fluid flow) conditions. (e) Capture efficiency in EHD-μD2 from 10× diluted blood spiked with prestained BT-474 cells (500 cells mL−1) under ac-EHD and pressure-driven flow (control; flow rate = 10 μL min−1) conditions. (f, g) Representative fluorescence images of prestained (DiD; red) BT-474 cells spiked in 10× diluted blood, nonspecific WBCs (CD45(+); green), and nuclear stain DAPI (blue) under ac-EHD (g, gˈ, gˈˈ) and pressure-driven flow (control; flow rate = 10 μL min−1) conditions (f, fˈ, fˈˈ) in EHDμD2. (h) Recovered cell numbers from PBS (black ■) and lysed blood (gray ●) spiked with prestained 50−1000 MCF7 cells mL−1 under ac-EHD in EHD-μD2. Data presented in (c−h) were obtained using the ac-EHD field strength of f = 1 kHz and Vpp = 100 mV. Each data point in (a−e) and (h) represents the average of three separate trials (n = 3), and error bars represent standard error of measurements within each experiment.
with earlier reports demonstrating fluid flow using an array of asymmetric microelectrodes energized by a single ac signal on the order of kHz and at low voltages (Vpp ∼1 V).7,8 In EHDμD2, it was observed that the presence of microtips facilitates the generation of complex fluid microvortices (video 4 for EHD and video 5 for control device in Figure S5, Supporting Information). This is believed to be due to the generation of nonuniform local forces on the microtip electrode surface that contribute to the net lateral flow resulting in fluid micromixing (Figure 2). Additionally, the microtips provide high aspect ratio structures with additional surface area and disrupt the streamlines to induce better fluid mixing thereby increasing the number of particle−electrode interactions. The critical parameters that influence fluid flow (e.g., surface shear force manipulation) and concomitant micromixing include the applied frequency and amplitude (along with electrolyte concentration and device geometry). To determine the optimal ac-EHD field strength for cell capture, capture efficiency of the devices was determined using whole cells expressing Human epidermal growth factor receptor 2 (HER2), an important biomarker and therapeutic target in breast cancer.14 To establish the fidelity of nanoshearing phenomenon in detecting low cell numbers, we chose breast cancer cell lines with variable levels of HER2 expression. Initially, we tested MCF7and T-47D cell lines for experiments with spiked PBS
ionic strengths used in this study (PBS, conductivity, Λ ∼ 15.4 mS/m), the electrical double layers are typically on the order of 1−2 nm12 in size, meaning this system engenders fluid flow within molecular distances of the electrode surface. We believe this is one of the most fascinating features of this phenomenon and have therefore termed the ability to tune fluid shear forces at the nanoscale as nanoshearing, given the capacity of these shear flows to displace weakly bound analytes on the surface. To investigate the nature of ac field induced fluid flow, fluorescent latex particles (diameter (d) = 970 nm; zeta potential (ζ) = −40 mV) were monitored using an upright microscope (Figure S4, Supporting Information). In EHDμD1, unidirectional flow was observed with the particles traveling from the smaller to larger electrode (see video 1 in Figure S4, Supporting Information), and no particle movement was observed in the absence of ac field (see video 2 in Figure S4, Supporting Information). We also measured the particle velocity using fluorescence latex particles (d = 960 nm) with zeta potential of −17 mV (see video 3 in Figure S4, Supporting Information). Under the experimental conditions used (frequency ( f) = 600 Hz; peak-to-peak amplitude (Vpp) = 0.1 V), almost similar particle velocity (30 ± 3 μms−1 in EHDμD1) was observed for both high and low charged particles. These observations indicate that the observed fluid flow could possibly be induced due to ac-EHD forces6,13 and corroborate 2045
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demonstrated by spiking prestained MCF7 along with HER2(-) MDA-MB-231 cells. When we spiked 200 MCF7 cells along with 105 MDA-MB-231 cells in PBS, a maximum of only 2% decrease in capture efficiency was observed (Figure 3d), indicating that our device can detect a small number of cells in the presence of a large excess of nontarget cells (video 6 in Figure S8, Supporting Information is a 3D animation demonstrating the micromixing effect in EHD-μD2 for specific cell capture). Additionally, a >8% reduction in capture efficiency was observed without fluid flow for mixed samples (MCF7 + MDA-MB-231 cells) in comparison to those obtained for only MCF7 cells (Figure 3d), presumably because of MDA-MB-231 cells being nonspecifically adsorbed on the sensing electrodes. A likely explanation for these results is that the forces generated due to ac-EHD field induced nanoshearing were large enough to shear away loosely adhered nonspecific cells from the antibody-coated surface. We then compared the capture efficiency of the device under different ac-EHD flow rates with those obtained under pressure-driven flow (via a syringe pump) of the same flow rates (Figure S9, Supporting Information). MCF7 cells (200 cells mL−1) were spiked in PBS and run on anti-HER2 functionalized devices under different ac-EHD flow rates obtained using the field strength of f = 600 to 100 kHz at constant Vpp = 100 mV. The equivalent flow rates calculated on the basis of the time required to flow 1 mL of blood under the ac-EHD field was 7.5−38.5 μL min−1, and these flow rates were used for pressure-driven flow experiments. Unlike ac-EHD case, the capture efficiency under the pressure-driven flow system was found to increase slowly at low flow rates (at 10 μL min−1, the capture efficiency decreases gradually with increasing flow rates (Figure S9, Supporting Information). However, a significant enhancement in capture efficiency across all operating ac-EHD flow rates was observed in comparison to pressure driven flows. This enhanced capture efficiency under ac-EHD induced fluid flow is presumably owing to the additional effective manipulation of shear forces (i.e., nanoshearing) and concomitant fluid mixing that can augment the specific capture of cells due to increased number of effective cell−surface (antibody functionalized) collisions. To validate the capture efficiency of our devices in complex heterogeneous samples, we spiked HER2(+) BT-474 cells in 1 mL of 10× diluted blood samples and performed experiments using EHD-μD2. Under the optimal ac field strength of f = 1 kHz and Vpp = 100 mV, the capture efficiency was found to be 87 ± 1.5% (Figure 3e). Control devices operated under a pressure driven flow rate of 10 μL min−1 resulted in capture efficiency of 63 ± 4.7%. An approximately 24% increase in capture efficiency with the use of ac-EHD induced fluid flow compared to pressure driven flow was achieved, which is attributed to additional nanoshearing and concomitant fluid micromixing. Representative fluorescence images for prestained BT-474 cells and nonspecific cells (i.e., CD45(+) WBCs) are shown in Figure 3f,g (bright-field and corresponding fluorescence images are also given in Figure S10, Supporting Information).
and lysed blood samples. Since reports on HER2 expression in MCF7 are inconsistent in the literature,15,16 immunoprecipitation followed by Western blotting was performed which verified the HER2 expression in our MCF7 or T-47D cell lines. We later tested our devices using BT-474 cells showing HER2 overexpression16 for experiments with spiked blood samples. For cell capture experiments (see Experimental Section), the samples were prepared by spiking prestained HER2(+) cells (e.g., MCF7 or T-47D) into PBS (10 mM, pH 7.0) and run on anti-HER2 functionalized devices (Figure 2 for cell capture and detection) under the frequency range of 600 Hz to 100 kHz at constant Vpp (100 mV). For both the devices, initial capture efficiency was found to increase sharply and then decrease gradually with increasing frequency (Figure 3a,b), presumably owing to ac-EHD induced fluid flow and manipulation of shear forces within the double layer (e.g., 1/ κ). Since the affinity interactions between cell and antibody are highly dependent on the flow rate, the resulting high capture efficiency at low frequency ranges is probably due to the stimulation of fluid flow around the sensors that can maximize the effective cell−antibody collisions (a condition where shear force < antibody−cell affinity force). In contrast, stronger fluid flow could possibly wash out target cells that can significantly reduce the effective antibody−antigen collisions/affinity interactions (a condition where shear forces > antibody−cell affinity force). Furthermore, stronger fluid flow can also induce the leaching of antibodies from the electrode surface. This can ultimately reduce the capture efficiency and affect the overall device performance. However, under the optimal applied field strength, the capture efficiency of our devices was highly reproducible with the relative standard deviation (% RSD) of 100 cells. We then spiked 50 to 1000 MCF7 cells in lysed rat blood (Figure 3h); capture efficiency of 48 ± 7.02% was attained for a seed level of 50 cells mL−1. It was also noted that the devices exhibited about less than 2% decrease in capture efficiency for lysed blood samples containing >100 cells mL−1 in comparison with buffer. These data indicate that the device performance was consistent for samples containing >100 cells mL−1 regardless of being spiked in buffer or lysed blood. We further tested the dynamic range of detection in blood samples by spiking BT-474 cells ranging from 50 to 1000 cells in 10× diluted blood. Under the optimal ac-EHD condition, the recovery for the seed level of 50 cells mL−1 was 53 ± 4.39% and capture efficiency was 86 ± 2.74% for samples containing >100 cells (Figure S12, Supporting Information). This level of capture efficiency obtained from lysed blood (e.g., 90 ± 1.7% for 100−1000 MCF7) and blood samples (e.g., 86 ± 2.74% for 100−1000 BT-474 cells in large excess of nonspecific cells including WBCs, RBCs, platelets, and other molecules) indicate that our method can be used for the efficient recovery of low cell numbers from complex heterogeneous samples without sample preprocessing. To demonstrate the utility of the nanoshearing phenomena in removing weakly bound cells present in blood samples, we performed experiments using 10× diluted blood samples in EHD-μD2 under the optimal ac-EHD field (e.g., f = 1 kHz, Vpp = 100 mV) in comparison with pressure driven fluid flow based 2047
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Foundation of Australia (CG-08-07 and CG-12-07) to M.T. These grants have significantly contributed to the environment to stimulate the research described here. The fabrication work was performed at Queensland node of the Australian National Fabrication Facility (Q-ANFF).
anti-HER2 functionalized devices under the optimal ac-EHD field. Trypan blue staining (Figure 4d) of the recovered cells resulted in >90% cell viability in EHD-μD2 (Figure 4e), suggesting that our approach can facilitate further molecular characterization of the recovered cells.
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CONCLUSIONS We have demonstrated a simple methodology for removing nonspecific adsorption of cellular species in 10× diluted blood. The key to the functionality and simplicity of our method lies in the innovative use of ac-EHD induced fluid flow for microand nanoscopic shear force manipulation (which we define as nanoshearing) to avoid nonspecific adsorption and enhance capture efficiency. This fuses with other innovative features of our platform such as: (i) fabrication of a long array of microelectrodes (sequential cathode and anode asymmetric pairs) within a long channel, which act both as fluid pumps (avoids use of additional pumping instruments) and capture domain, (ii) capability of analyzing large sample volume (highly desirable for cancer cell isolation in clinical samples), and (iii) noncovalent binding of antibodies (avoids the need for using covalent coupling chemistries). For microtip electrodes, these provide a very sensitive methodology to control ac-EHD flow vortices around the microtip to specifically capture target cells in large excess of nontarget cells and other molecules. The analytical performances of our method (87 ± 1.5% capture efficiency for 500 cells mL−1 in blood, with a 4-fold decrease in nonspecific adsorption of blood cells) suggest that ac-EHD devices will be useful for the quantification of low-abundance cancer cells from complex body fluids. Similar to the microtip electrodes, a change in aspect ratio of the planar electrodes can result in additional asymmetry in the electrode structure, which can also produce more complex fluid vortices. Therefore, acEHD devices with high aspect ratio electrode structures can also be beneficial to achieve improved analytical performance of the devices. We further envision that this proof-of-concept demonstration is capable of stand-alone use or can be used in tandem with current technologies for cell capture.17−25 The potential to preserve cell viability could facilitate detailed molecular and functional characterization of cells making our approach potentially attractive for numerous biological and medical applications.
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ASSOCIATED CONTENT
S Supporting Information *
Design, fabrication, and characterization of the ac-EHD devices and additional data. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. Tel: +61-7-33464178. Fax: +61-7-33463973. *E-mail:
[email protected]. Tel: +61-7-33463973. Fax: +61-733463973. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the ARC DECRA (DE120102503) to M.J.A.S. We also acknowledge funding received by our laboratory from the National Breast Cancer 2048
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