Microfluid Nanofluid (2012) 13:613–623 DOI 10.1007/s10404-012-0978-7
RESEARCH PAPER
Continuous-flow in-droplet magnetic particle separation in a droplet-based microfluidic platform Hun Lee • Linfeng Xu • Byungwook Ahn Kangsun Lee • Kwang W. Oh
•
Received: 25 January 2012 / Accepted: 27 March 2012 / Published online: 20 April 2012 Ó Springer-Verlag 2012
Abstract This paper presents a continuous-flow in-droplet magnetic particle separation in a droplet-based microfluidic device for magnetic bead-based bioassays. Two functions, electrocoalescence and magnetic particle manipulation, are performed in this device. A pair of charging metallic needles is inserted into two aqueous channels of the device. By electrostatic force, two different solutions can be merged to be mixed at a junction of droplet generation. The manipulation of magnetic particles is achieved using an externally applied magnetic field. The magnetic particles are separated by the magnetic field to one side of the droplet and extracted by splitting the droplet into two daughter droplets: one contains the majority of the magnetic particles and the other is almost devoid of magnetic particles. The applicability of the continuous-flow in-droplet magnetic particle separation is demonstrated by performing a proof-of-concept immunoassay between streptavidin-coated magnetic beads and biotin labelled with fluorescence. This approach will be useful for various biological and chemical analyses and compartmentalization of small samples. Keywords Magnetic particle manipulation Magnetic field Magnetic bead assay Droplet merging Droplet splitting
H. Lee L. Xu B. Ahn K. Lee K. W. Oh (&) SMALL (Sensors and MicroActuators Learning Laboratory), Department of Electrical Engineering, University at Buffalo, The State University of New York (SUNY at Buffalo), Buffalo, NY 14260, USA e-mail:
[email protected]
1 Introduction Droplet-based microfluidics is a promising microfluidic platform for high throughput, continuous flow, and ultralow volume studies of biological and chemical reactions. One of the key unit functions to perform reactions within individual droplets is precise manipulation of pairwise droplets for droplet coalescence. In the case of passive coalescence, some geometries of channel were used to control the location of droplet coalescence (Tan et al. 2004, 2007). Proper coalescence still poses challenges in many cases since it depends on droplet frequency matching under high throughput conditions. In general, pairwise droplets should be precisely synchronized in space and time (Ahn et al. 2006, 2011). Droplet coalescence has been straightforwardly demonstrated for droplets that are not treated by surfactant (Ismagilov 2003; Zheng and Ismagilov 2005). However, it is difficult to coalesce droplets when they are stabilized with surfactant, which needs more complex additional processing. In the case of active coalescence, several strategies have been used to realize droplet coalescence by externally applied forces, such as electrical, (Schwartz et al. 2004; Zagnoni and Cooper 2009), heatbased (Kohler et al. 2004), or optical methods (Lorenz et al. 2006). One of the widely utilized methods is the use of electric fields at a region where pairwise droplets meet. In this configuration, oppositely charged droplets were generated by the simultaneous application of voltages and merged by their charges, causing electrostatic attractive forces between them. As the droplets get closer, the electric forces can be amplified by dipole–dipole interactions between the droplets, resulting in destabilization of the surface of the droplets to be merged (Gu et al. 2011). However, in this approach the charged pairwise droplets still need to be synchronized at the location where electrocoalescence occurs.
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The difficulty of merging pairwise droplets at arbitrary points has limited the widespread use of droplet-based microfluidic devices for various biological applications. Microfluidic immunomagnetic systems have been demonstrated due to ease of manipulation in a fluidic environment using external magnetic fields (Fan et al. 1999; Jemere et al. 2002; Nolan and Sklar 2002; Choi et al. 2002). The systems use biochemically functionalized magnetic particles that bind to specific targets to sort the magnetic particles from a solution by deflecting them (Kim and Park 2005). This technology is advantageous in that the magnetic particle has a high surface-to-volume ratio, and thus it can provide relatively large binding sites for biochemical reactions. It enables multiplexed assay by using differently functionalized magnetic particles, enhanced sensitivity due to the large surface area per unit volume, and short reaction time due to the freely mobile suspensions in fluids (Peyman et al. 2009). The manipulation of the magnetic particles can be achieved without complex or expensive equipment to handle them in a medium of microfluidic channel. A simple permanent magnet is enough to induce a sufficient field gradient throughout a channel to move or trap them (RodriguezVillareal et al. 2011). Recently, an electromagnet on the microfluidic system was reported to manipulate magnetic particles (Siegel et al. 2006). It is advantageous because it can be switched on and off quickly using electrical signals and the strength of magnetic field can be adjusted. For these reasons, the magnetic particles have been used for DNA separation, protein digestion, immunoassays, and cell capture (Jeong et al. 2008; Wang et al. 2000; Auroux et al. 2002; Beebe et al. 2002). However, the concept still requires a batch procedure with a fixed sample volume. Another conventional separation method is to deflect magnetic particles from their continuous flow path by an externally applied magnetic field (Pamme 2007). The continuous flow handling of magnetic particles has been utilized as a technique for separating magnetic particles and cells from a sample stream (Blankenstein and Larsen 1998; Kim and Park 2005; Pamme and Manz 2004; Pamme and Wilhelm 2006; Siegel et al. 2006; Yung et al. 2009). An advanced continuous flow procedure has recently been demonstrated by traversing functionalized magnetic beads across multiple co-laminar flow streams (Peyman et al. 2009). The magnetic particles were deflected through multi-laminar reagent streams and consecutive binding and washing steps were performed on their surface. Despite its huge potential, the continuous flow magnetic handling has so far suffered from the lack of re-configurability and compartmentalization for isolated magnetic bead-based bioassays. Its ability to increase reagent incubation periods would be limited. The compartmentalization of reagents within droplets is an extremely effective way to prevent contamination and dilution effects between isolated multiple bioassays and increase the
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incubation time of samples by lengthening the channels or by collecting the droplets in containers. The droplet-based platform has potential for high throughput and ultralow volume studies of various forms of chemical synthesis and biological analysis. Recently, Lombardi and Dittrich (2011) have reported a concept of magnetic particle separation in a droplet by splitting the droplet into two to investigate drug–protein interactions, which is similar to our system in the magnetic particle separation. However, the system does not have a function to merge droplets with different reagents to extract a target reagent within the droplets and allow mixing between reagents on the chip. Thus, it is necessary to carry out an off-chip procedure to mix them together. In this work, we demonstrate continuous flow in-droplet magnetic particle separation in a droplet-based microfluidic device for magnetic bead-based bioassays. The device consists of two parts: one is for electrocoalescence and the other is for magnetic particle manipulation to carry out bead-based streptavidin–biotin reactions within the droplets. The electrocoalescence is successfully achieved by applying electric fields without any accidental coalescence between the droplets with different solution at a region of droplet formation. In terms of the electrocoalescence, the key idea of our device is that this method is simple and robust compared with other electric coalescence methods because it enables more reliable coalescence without any precise synchronization in spacing and time between the droplets. Furthermore, it does not require precise electrode alignment and a metal deposition for the electrode. First, two solution streams charged by metallic needles are polarized; then the applied electric fields induce the attraction forces between them at the location of droplet generation where the merging occurs. The device enables ondemand droplet coalescence by switching on and off a voltage source. Furthermore, the electrocoalescence has the potential to merge several droplets by integrating electrodes where the coalescence occurs for a post processing. We experimentally demonstrate the in-droplet manipulation and separation of the functionalized magnetic beads by the externally applied magnetic field within each merged droplet. Subsequently, the droplets are split into two daughter droplets at a T-junction to separate the portion of the droplet that contains the majority of the magnetic beads. This approach enables the continuous-flow in-droplet magnetic bead separation in the droplet-based microfluidics.
2 Design and fabrication 2.1 Design This work demonstrates a magnetic bead-based bioassay in a droplet-based continuous-flow microfluidic device
Microfluid Nanofluid (2012) 13:613–623 Fig. 1 Schematic illustration of the proposed device. a the working principle, b the droplet generation module, and c splitting droplets into two, with magnetic beads and without magnetic beads, at T-junction
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(a)
(b)
(Fig. 1). A flow-focusing T-junction where an island structure is configured at the centre of the main channel was used to ensure the generation of monodispersed droplets in a robust manner. In order to charge the water phases 1 (i.e., suspension of magnetic beads coated with streptavidin) and 2 (i.e., suspension of biotin with fluorescence label), the metallic needles were inserted into each of the access holes behind inlet 1 and inlet 2. The channel between the access holes and inlets was filled with water to electrically connect them. The immiscible phase (i.e., mineral oil) was injected into the middle inlet to generate the droplets at the flowfocusing junction. At the location where the electrocoalescence occurs, the circular shape of the channel was configured to decrease the pressure caused by the water phases pushing the oil phase back, which can cause unstable droplet formation and electrocoalescence. Subsequently, for mixing the two water phases together, U-shape channels were configured, thereby confirming the reaction between the magnetic beads coated with streptavidin and fluorescently labelled biotin after the electrocoalescence. For more effective mixing between reagents in the droplet, (Bringer et al. 2004) suggested a meander-shape channel with short turns. However, in this device, U-shape channels with a long path were used to increase the incubation time of the magnetic beads in the droplets, since the incubation time can be one of the major parameters in determining the fluorescence intensity of beads and assay sensitivity. A permanent magnet was positioned at an adjacent channel, so that it generated a
(c)
magnetic field perpendicular to the channel. The magnetic field was then introduced to pull the magnetic beads towards one side within the droplets. The droplets were then each split into two droplets, one containing the majority of the magnetic beads collected at outlet 1, and the other collected at outlet 2. 2.2 Microchip fabrication Our device was fabricated using conventional soft-lithography techniques (Xia and Whitesides 1998). The 3-in. silicon wafer (University Wafers, South Boston, MA, USA) was submerged into buffered hydrofluoric acid (BHF) at room temperature for 5 min to remove a thin oxide layer that can cause poor adhesion between the SU-8 and the surface of the wafer. Afterwards, it was cleaned with acetone, followed by methanol. It was then rinsed with deionized water and blown dry with filtered nitrogen gas. The cleaned wafer was placed on a hot plate at 120 °C for 5 min for complete dehydration. The SU-8 (SU-8 2050, Micro-Chem Corp, Newton, MA, USA) was then spin-coated with the target thickness (i.e., 50 lm) on the cleaned wafer using a spin processer (WS-650Mz NPP from Laurell Technologies, North Wales, PA, USA). After the spin-coating process, soft bake was performed on a hot plate for 3 and 9 min at 65 and 95 °C, respectively. UV photolithography was carried out using a contact mask aligner. Post exposure bake (PEB) was conducted on the
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hot plate for 2 and 7 min at 65 and 95 °C, respectively, followed by development in the SU-8 developer for 5 min. After the development, it was washed with isopropyl alcohol. It was then blown dry with filtered nitrogen gas and placed on the hot plate at 100 °C for 5 min to evaporate the rest of the isopropyl alcohol. The measured thickness of the SU-8 pattern was 47 ± 2 lm. A prepolymer of PDMS (Sylgard 184, Dow Corning) and curing agent was thoroughly mixed at a ratio of 10:1 (wt/wt). The mixed PDMS was degassed in a vacuum chamber to remove air bubbles for 20 min. To easily peel off the PDMS from the SU-8 pattern, hexamethyldisilazane (Sigma Aldrich, Saint Louis, MO, USA) was silanized on the SU-8 master in the vacuum chamber at room temperature for 30 min. The permanent magnet for the magnetic particle manipulation was placed directly on the SU-8 pattern which was preformed about 1 mm away from the channel for defining the location of the magnet, and it was then fixed with a commercially available double-sided tape on the SU-8 pattern. The PDMS was carefully poured onto the SU-8 master mould and cured at 65 °C for 30 min. After curing, the PDMS replica was peeled off, defining the position of the magnet. Finally, the replicated PDMS was bonded irreversibly with a glass slide by exposing O2 plasma on the surface of the PDMS and glass slide. 2.3 Materials and methods The voltage was supplied by a high-voltage source (HVS448-1500, Lab-Smith, Livermore, CA, USA) connected to a computer controller to charge the two water phases. The magnetic fields were generated by a rectangular neodymium–iron–boron magnet (12.7 9 3.2 9 3.2 mm3, Emovendo Magnets & Elements, Petersburg, WV, USA). The mineral oil (M8410, Sigma-Aldrich) was used as the immiscible phase and 2 % nonionic surfactant (span-80, S6760, Sigma-Aldrich) was added to the mineral oil to prevent accidental coalescence. The channels were treated with a commercial surface coating agent (Aquapel, 47100, Pittsburgh Glass Works, PA, USA) to enhance hydrophobicity. The viscosity of the mineral oil is 34.5 cSt (Ward et al. 2005) and interfacial tension is estimated to be 49.255 mN/m from published data for mineral oil and water (Cohen et al. 2010). The concentration of fluorescently labelled biotin (biotin-4-fluorescein, Molecular Probes, Invitrogen) was 12.5 lg/ml. The magnetic beads (2.8 lm diameter, Dynabeads M-270 streptavidin, Invitrogen) were diluted in deionized water. The fabricated microfluidic device was connected through a silicone tube to a syringe pump (KDS100W, Fisher Scientific, IL, USA). All experimental results were captured by a CCD camera mounted on a Nikon stereo-type microscope.
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2.4 Electrocoalescence By incorporating the electrostatic forces in the flowfocusing T-junction geometry, the two water phases were merged into single droplets. This produces the robust electrocoalescence of droplets at the region where the water phases meet. The droplet size was controlled by the volumetric flow rates of the oil (Qoil) and the water (Qwater) at the T-junction structure. The control to merge the water phases is carried out by switching on and off the highvoltage source, which enables more reliable continuous flow generation of merged droplets. Figure 2 shows the equivalent electric circuit diagrams and photographs before and after electrocoalescence. The module for the electrocoalescence can be modelled as an electric circuit. The water phase behaves as a conductor with low resistance (Rwater), whereas the oil phase behaves as an insulator with high resistance (Roil). Since the two water phases are connected to different potentials (DV), electric fields occur between the two tips of the water phase separated by the oil phase; this will create a sort of capacitor (C). Before the merging of droplets, the water streams start to come out with opposite charges (i.e., positive and negative charges), by the injection flow from the syringe pump (Fig. 2a). The electric field between the tips of the water streams can be amplified as they get closer, which can lead to destabilization of the tip surfaces of the water streams (Priest et al. 2006), resulting in electrocoalescence by the attractive electrostatic force (Fig. 2b). The following oil stream pinches off the merged water streams, resulting in generation of the merged droplets with the injection flow. Once droplets are released (i.e., discharging of the capacitor), the water streams carry opposite charges with them (i.e., charging of the capacitor). 2.5 Magnetic field The magnetic fields were used to attract the magnetic beads towards one side of each droplet. The magnetic force, Fmag, on the magnetic beads is calculated by Fmag ¼
1 VDv rB2 2 l0
ð1Þ
where Dv is the net magnetic susceptibility of the beads in a medium, V is the volume of the beads, l0 is the permeability of free space, and B is the magnetic field (Zborowski et al. 1995). Another dominant force acting on them is viscous drag force, Fdrag, which is generated against the opposite direction of the moving magnetic beads (Tsai et al. 2011). Fdrag is represented by the following equation:
Microfluid Nanofluid (2012) 13:613–623 Fig. 2 Equivalent circuit diagrams a before merging b after merging by the applied voltage at the given flow rates. The length of scale bar is 100 lm
Fdrag ¼ 6prgm
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(a)
(b)
ð2Þ
where r is the radius of the beads, g is the viscosity of the medium, and m is the velocity of the beads. While being deflected by the magnetic fields, the magnetic beads also experience an equal force by the movement of beads, and thus the sum of Fmag and Fdrag equals zero: Fmag þ Fdrag ¼ 0
ð3Þ
Equation 3 gives us the velocity of the magnetic beads in the medium, which is calculated by m¼
VDv rB2 12prgl0
ð4Þ
We conducted a two-dimensional simulation with COMSOL Multiphysics 3.5a to show a magnetic field distribution from the permanent magnet used in our experiment. For the simulation, a magneto-statics module
was chosen and the magnetic flux density was obtained as a function of distance from the magnet based on the simulation result as shown in Fig. 3a. With the increasing distance away from the magnet, the magnetic fields are exponentially decreased. The velocity of the magnetic beads in the droplets was calculated with the assumption that the magnetic field is applied in the x-direction, perpendicular to the direction of flow. The magnetic beads move in a laminar flow as shown in Fig. 3b. The results show that the velocity of the magnetic beads is decided by the magnetic field gradient. With increasing distance away from the magnet, the magnetic fields are exponentially decreased. Equation 4, derived from the balance of forces, qualitatively shows what is expected experimentally. A convection of internal flow in the droplets is neglected with the assumption that the internal flow is almost perpendicular to the magnetic field (Kinoshita et al. 2007). We also chose to neglect a spatial bead position in the droplets with the fact that the width of
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618 Fig. 3 a Magnetic flux density as a function of distance away from the magnet. Inset shows the magnetic field distribution. b Velocity of magnetic beads in droplet
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(a)
the microfluidic channel used is 100 lm and the permanent magnet is located several millimeters away from the channel, which means that the variation of the beads’ position is negligible. However, it is noted that once the beads are on the magnetic fields, they can be magnetized with dipoles. The dipoles will tend to align from pole to pole in response to the externally applied magnetic field, resulting in an increased volume of beads, which is irregular motion. Thus, the velocity of the beads will increase significantly more than what we expect. For that reason, the separation of beads is empirically investigated. We expect that the velocity of the flow would typically be a more dominant parameter than the internal flow since when the velocity of the flow is increased, the residence time of the beads in the magnetic fields decreases. The beads will move less towards one side of the droplets as the flow velocity is increased. Therefore, we empirically investigated the separation working zone by varying the magnet position away from the channel at given total flow rate (Qwater ? Qoil) with corresponding to the velocity of the droplet in Fig. 4b. 3 Results and discussion First, we demonstrated that the two water streams were electrostatically charged by applying the potential difference through the metallic needles inserted into the access holes, followed by merging the two streams to generate droplets. The water and oil flow rates were set at 30 and 50 ll/h, respectively, and the corresponding droplet diameter was 125–130 lm. While the droplets were generating, DC voltage (20 V) was applied, which was sufficient to ensure the electrocoalescence of all droplets. This system can be considered as equivalent to an electric circuit, as shown in Fig. 2a. Rwater is the resistance due to the water stream between the electrode and the tip of the water stream, and Roil is the resistance caused by the oil stream between the two tips of the water stream. The two tips are roughly equivalent to a capacitor, C, which is repeatedly discharged when a droplet is released. The total
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(b)
voltage drop across these elements is the applied voltage, DVDC. Therefore, we get the following equation: DVDC ¼ 2Rwater I þ U
ð5Þ
where I is the current flowing through this system and U is the voltage at the capacitor. When the water streams are growing to produce the droplet, using Eq. 5 the charge (q) induced at the tip surface of the water stream is given by following equation: q ¼ C ðDVDC 2Rwater IÞ
ð6Þ
With the assumption that Rwater Roil and I is extremely small, Eq. 6 can be simplified by q ¼ CDVDC
ð7Þ
If we assume that when the droplet is released, the capacitor is completely discharged and all the charges are going into the droplets, the induced charges should be equal to CDVDC. The assumption is reasonable because water is conductive; there is no way to stay anywhere else other than on the end surface of the water stream. It is noted that it was impossible to estimate the current across this system to obtain the capacitance experimentally due to ambient noise since the current was extremely small. Instead, the charges induced can be calculated by balancing forces in which the capacitance can then be obtained. There are balancing forces acting on the droplet merging mechanism: the electrostatic force (Fe) that is accelerating to meet the two water streams and the viscous drag force of the oil (Fv) that is decelerating to prevent them from merging. Fe and Fv in our experiments are given by Fe ¼ qE;Fv ¼ 6pagDt
ð8Þ
where q is the charge per the tip of the water stream, E the electric field between the tips of the water streams, a the diameter of the tip of the water stream, g the viscosity of the mineral oil and Dt the relative velocity of the tip of the water stream to the oil, which was calculated by measuring the velocity of the water stream with voltage and without.
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(a)
(b)
Fig. 4 a Images of separation and no-separation condition by the magnetic field. b The distance to separate magnetic beads in droplets at different total flow rates and the velocity of droplet corresponding to the total flow rate. The scale is 100 lm
The sum of these should be zero: qE 6pagDt ¼ 0 From Eq. 6, we obtain rffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi 6pagDtr 2 er q¼ k
ð9Þ
ð10Þ
where k is coulomb constant, er is the relative permittivity of the oil medium, and r the distance between the tips of the water streams. The value r is initially formed by the structure in the middle of the channel. In this calculation, the minimum charge for the electrocoalescence induced by the applied voltage is equal to 6.6 9 10-14 C when a = 60 lm, g = 28.1 Ns/m2, Dt = 6 22.5 lm/s, r = 30 lm, er = 2.2 and k = 9 9 109 Nm2/C2. Based on
the charges calculated by Eq. 10, we obtain the minimum voltage, 9.0 V; E is proportional to the voltage applied and inversely proportional to the distance between electrodes based on the assumption that the fields are uniform. From Eq. 7, the capacitance is equal to 7.3 9 10-15 F. But, when we applied 10 V, the droplet generation was unstable. To ensure successful electrocoalescence, we increased the voltage to 20 V. When we applied a voltage of 20 V, the two water streams could be merged at the junction, generating the droplets. The droplets were generated at a rate of 10 Hz. Qualitatively speaking, it is clear that the electric field will attract the end surfaces of the water streams to each other and can be increased by dipole– dipole interaction as they get closer. We believe that the electrostatic force can lead to instability of the end surfaces
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and so break the interface between the water and the oil. From a different point of view, considering the balancing forces acting on the droplet merging mechanism, an oil film resistance can be also considered instead of the viscous drag force since the electrostatic force between the tips of the two water streams is inducing an attractive force, squeezing the oil film out of the way. For a clear analysis, it is necessary to study more deeply the mechanism of the electrocoalescence for an accurate theoretical model. Once we turned off the voltage, there was no merging between the two streams and droplet pairs were generated. The response time was \0.24 s. This was measured by subtracting the last time when the merged droplets were generated from the start time when the droplet pairs were generated under the assumption that once the voltage source is turned off, there are no induced charges from it. We investigated the separation working zone at the channel region being affected by the magnetic field. The magnet position was varied away from the channel from 1 to 5 mm, and the total flow rate (Qwater ? Qoil) was varied up to 500 ll/h while the flow rate ratio (Qratio = Qoil/Qwater) was fixed at 2. As shown in Fig. 4, we categorized the separation condition into two zones: separation and no separation. In the separation zone, the magnetic beads are moved and directed to one side within the droplets by the applied magnetic field. In the no separation zone, the magnetic beads are randomly distributed in the droplets at the given magnetic fields. The residence time of the magnetic beads in the magnetic field influences the separation efficiency of the beads within the droplets. For the magnetic bead manipulation, the permanent magnet was directly placed about 1 mm away from the channel and the magnetic field strength corresponding to the location of the magnet was 319.68 mT. The arrows in Fig. 4a indicate the relative magnetic field strength required to separate the magnetic beads towards one side of the droplet. Thus, by varying the magnet position and total flow rate, we obtained the result as shown in Fig. 4b. When the total flow rate increased, the residence time decreased in the magnetic fields, resulting in no separation. With increasing distance between the magnet and channel, the magnetic fields exponentially began to decrease. To demonstrate the utility of this device for biological applications, a well-established biochemical reaction (i.e., biotin–streptavidin binding assay involving a single binding–separation step) was performed using the magnetic beads coated with streptavidin and biotin labelled fluorescently. We first investigated the off-chip binding condition. It is necessary to remove excess biotin that can increase background noise when we detect fluorescence intensity after the reaction in droplets. 10 ll of streptavidincoated magnetic beads with *5 9 108 /ml concentration was mixed with different fluorescently labelled biotin
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concentrations (e.g., 25, 12.5, 6.25, and 3.12 lg/ml). The mixtures were placed on a microscope slide and measured under the stereo-type microscope to obtain fluorescence images. We chose a 3.12-lg/ml concentration of biotin that enabled a relatively clearer fluorescence image (image not shown). Once the bioassay had been successfully performed offchip, the procedure was adapted for on-chip continuousflow in-droplet magnetic beads separation, as shown in Fig. 5. The electrocoalescence was successfully achieved for on-demand formations of the droplets at the given flow rate, Qwater = 30 ll/h and Qoil = 60 ll/h and the droplets were generated at a rate of 10 Hz. The concentration of the biotin used in the microfluidic device was 3.12 lg/ml. Based on the information provided by the manufacturer, we can calculate the biotin-binding capacity of magnetic beads corresponding to their concentration (*5 9 108/ml). The calculated capacity of biotin concentration was 0.95 lg/ ml, so we expect that there is potential for this system to detect lower biotin concentrations. However, our detection system was limited for lower biotin concentrations. To enhance the detection limits and to increase the sensitivity of assay, a more sophisticated CCD camera should be used. Before injecting the magnetic beads into the inlet, they were agitated by shaking manually in a microcentrifuge tube to randomly distribute them. The water and oil phase flow rates were set at 30 and 60 ll/h, respectively, and the corresponding droplet size was 120–125 lm. The droplets were generated by the electrocoalescence to mix them together. The droplet generation was significantly stable at the given flow rate by adding the surfactant in the oil phase and coating the channels with the commercial coating agent to increase hydrophobicity. The droplets containing the magnetic beads and biotin solution were subsequently passed through the mixing channel (100 lm width). Before entering the magnetic field generation part, the magnetic beads were randomly moved in the droplets. For the magnetic bead manipulation, the permanent magnet was directly placed about 1 mm away from the channel. As the droplets with a plug shape moved downstream, the magnetic beads began to be separated to one side within droplets according to stronger magnetic field gradients. Once concentrated, the magnetic beads were split into two droplets equally, flowing out of the two branched channels (80 lm width). The droplets containing the majority of the magnetic beads were subsequently collected from outlet 1 and the droplets not having them were collected from outlet 2, as shown in Fig. 6. This system does not have the ability to perform washing steps to reduce the background noise caused by excess reagents for multi-step reactions. However, to help overcome this limitation, a post processing to dilute the reagent in the droplets can be used. After the collection of the split droplets, the droplets can be
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Fig. 5 Experimental results for continuous-flow in-droplet magnetic beads separation in a droplet-based microfluidic device. The scale is 200 lm
re-injected into another device where merging between the reagent and washing droplets can be performed by the electrocoalescence. The incubation time of magnetic beads in droplets at the given flow rate was approximately *31 s. It is noted that undesired merging of droplets occurred in the outlets because the more droplets are
collected in the outlets, the more the droplets touch others in their vicinity. In order to prevent this, a minimum distance should be kept between droplets, or different surfactants can be used to stabilize the droplets. We analysed the error rate defined as the ratio of the number of droplets with magnetic beads to the number without, by counting
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(a)
(b)
Fig. 6 Photographs of fluorescent magnetic beads. Original images captured by fluorescent microscope were darker than these images, but the fluorescent intensity was increased to show the clear magnetic
beads in the droplets. a The magnetic beads combined with fluorescently labelled biotin in the droplets b no magnetic beads in the droplets. The circles indicate the separation errors. The scale is 200 lm
the number of droplets with magnetic beads at outlet 2. The error rate obtained was below 0.5 % when the magnet was located 1 mm away from the channel with Qwater = 30 ll/h, Qoil = 60 ll/h and a bead concentration of *5 9 108/ml. For highly sensitive studies of immunoassay, it is important to maintain a consistent magnetic bead concentration within each droplet. In order to prove the magnetic bead concentration, the droplets containing magnetic beads can be collected in a tube. Subsequently, the droplets are placed between two cover slips to make all of the beads visible in one focus plane. It is then possible to analyse the beads using the images collected with the CCD camera. The number of beads can be counted using a particle analyser. As a future study, we will try to investigate the concentration of magnetic beads further to achieve high accuracy and ensure sensitivity. This demonstration will help when assessing the platform’s applicability for generic magnetic separation procedures with more than one binding step (i.e., sandwich immunoassay involving two bindingseparation steps) with a minor modification of the device design, which is under investigation and will be reported later. This concept-of-principle demonstrates the potential applicability of the droplet-based microfluidic platform technology for high throughput, continuous-flow, and magnetic bead-based immunoassays.
formation. We have shown that the manipulation of magnetic beads could be easily controlled by the magnetic fields from the simple permanent magnet for in-droplet separation. By varying the surface functionality of the magnetic beads, the proposed method could be used to perform many useful and analytical bead-based immunoassays in the continuous-flow droplet-based microfluidic platform.
4 Conclusion We have successfully demonstrated continuous-flow in-droplet magnetic particle separation in the droplet-based microfluidic device for magnetic bead-based bioassays. Robust electrocoalescence was achieved by electrostatic force without any accidental coalescence between the droplets with different solution at the region of droplet
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Acknowledgments This work was supported by the National Science Foundation grants (Grant Nos. ECCS-1002255 and ECCS-0736501).
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