We thank the Kenis group at the University of Illinois at Urbana-Champaign for helpful discussions and generously providing use of cleanroom facilities.
This work was funded by an NIH Pathway to Independence PI Award, under Grant No. 4R00HG004183-03 (Charles M. Schroeder and Melikhan Tanyeri).
This work was supported by the National Science Foundation through a Graduate Research Fellowship to Eric M. Johnson-Chavarria.

The hydrodynamic trap consists of a two-layer hybrid (polydimethylsiloxane (PDMS) /glass) microfluidic device for particle confinement. Steps 1-2 describe fabrication of microfluidic devices, and Steps 3-4 discuss device design and operation.
1. SU-8 Mold Fabrication (not shown in video)
<ol>
	<li>Clean two silicon wafers (3" diameter) with acetone and isopropyl alcohol (IPA).</li>
	<li>Dry wafers with N2 and place them on a hotplate at 65&deg;C for 1 min to remove residual moisture.</li>
	<li>Spin coat wafer #1 with SU-8 2050 photoresist (PR) for 30 sec at 4000 rpm to create a ~40 &mu;m thick mold for the fluidic layer. Spin coat wafer #2 with PR for 30 sec at 1500 rpm to create a ~150 &mu;m thick mold for the control layer.</li>
	<li>Soft bake wafer #1 at 65&deg;C for 3 min and then at 95&deg;C for 6 min. Soft bake wafer #2 at 65&deg;C for 5 min and then at 95&deg;C for 20 min.</li>
	<li>Expose wafers to UV with their respective masks (wafer #1: ports and fluidic channels, wafer #2: port and control layer) and appropriate exposure intensity (~150 mJ/cm2, ~260 mJ/cm2 respectively).</li>
	<li>Post bake wafer #1 at 65&deg;C for 1 min and then at 95&deg;C for 6 min. Post bake wafer #2 at 65&deg;C for 5 min and then at 95&deg;C for 10 min.</li>
	<li>Develop wafers with propylene glycol methyl ether acetate (PGMEA) until uncured PR is removed. Rinse wafers with IPA and dry with N2.</li>
</ol>
2. Microfluidic Device Fabrication
<ol>
	<li>Silanize the surface of the SU-8 molds by placing the wafers in a desiccator under vacuum for ~10 min with a glass dish containing a few drops of trichlorosilane. Surface silanization assists in peeling the (PDMS) replicas off the SU-8 molds.</li>
	<li>Mix and degas PDMS in base:crosslinker ratios of 15:1 and 5:1 for the fluidic and control layers respectively.</li>
	<li>Spin coat the 15:1 PDMS mixture onto the fluidic layer mold (wafer #1) for 30 sec at 750 rpm and then place the wafer into a Petri dish. Place the control layer mold into a Petri dish and pour 5:1 PDMS mixture onto the mold to a thickness of ~ 4 mm.</li>
	<li>Bake wafers/PDMS for 30 min at 70&deg;C to partially cure the PDMS layers.</li>
	<li>After cooling down the wafers/PDMS to room temperature, cut the PDMS replica, which will form the control layer (wafer #2), from the Petri dish with a scalpel and peel it off the SU-8 mold. Hole punch an access port to the microchannel that will act as the on-chip membrane valve with a 21 gauge needle.</li>
	<li>Place the PDMS replica with the control layer onto the wafer #1 (which has the spin-coated PDMS fluidic layer). Carefully align and seal the control layer to the fluidic layer using a stereo microscope. Make sure to remove all air pockets between the layers and bake at 70&deg;C overnight to fully cure both layers. This baking step will result in a monolithic PDMS slab with two layers.</li>
	<li>After cooling to room temperature, cut and peel the PDMS replica containing both the control and the fluidic layers off the SU-8 mold using a scalpel. Remove excess PDMS and separate each device unit with a razor blade. Hole punch access ports to the microchannels in the fluidic layer with a 21 gauge needle.</li>
	<li>Bond the PDMS slab to a coverslip to obtain a complete device. First, clean a coverslip (No: 1.5, 24 x 45 mm) with acetone and IPA. Next, treat both the coverslip and the PDMS replica surfaces with oxygen plasma under 500 mTorr for 30 sec, and immediately bring the two surfaces into contact to form an irreversible seal.</li>
	<li>Bake the devices overnight to increase bonding between the PDMS layers and the coverslip.</li>
</ol>
Steps 3-4 describe implementing the hydrodynamic trap using the microfluidic device described above.
3. Hydrodynamic Trap Experimental Setup
<ol>
	<li>Place the microfluidic device onto the stage of an inverted microscope and secure it with stage clips.</li>
	<li>Fill two gas-tight syringes separately with buffer and sample solutions and place them on a Harvard Apparatus Syringe pump (PHD 2000 Programmable). The buffer and the sample solutions are delivered to the microfluidic device through a 1 ml and a 250 &mu;l syringe, respectively. Typically, a 50 mM Tris/Tris HCl buffer solution (pH 8.0) containing 0.02% v/v Triton X-100 is used as the buffer solution. The sample solution consists of a particle suspension (e.g. fluorescent polystyrene beads) in the buffer solution.</li>
	<li>Establish the fluidic connections between the syringes (delivering the sample and the buffer) and the microfluidic device. Connect the syringes to 1/16" outer diameter (OD) x 0.020" inner diameter (ID) perfluoroalkoxy (PFA) tubing using luer-lock adapters. Connect the other end of the PFA tubing to the inlet ports of the microfluidic device with 24 gauge metal tubing. A T-valve can be placed between the sample syringe and the sample port on the microfluidic device to control sample delivery.</li>
	<li>Establish the fluidic connections for the outlet channels in the microfluidic device. Connect the two outlet channels to PFA tubing (1/16" OD x 0.020" ID) by using 24 gauge metal tubing. The PFA tubing for the outlets should be of equal length. Submerge both outlet tubes into a 1.5 mL centrifuge tube filled with buffer solution, which serves to maintain a constant pressure drop between the syringes and the outlet channels.</li>
	<li>Fill the on-chip valve with fluorinated carrier oil using a 3 mL luer-lock plastic syringe to prevent air from leaking into the fluidic layer during operation. The air in the valve chamber is pushed through the PDMS membrane into the microchannel in the fluidic layer and later removed from the device with fluid flow through the outlet ports.</li>
	<li>Connect a pressurized inert gas (nitrogen) supply to the port in the control layer for on-chip valve operation. For this purpose, we use a nitrogen tank (2200 psi) and an electronic pressure regulator to supply 0-30 psi to the on-chip valve in the microfluidic device. The nitrogen tank is connected to the pressure regulator using &frac14;" OD x 0.170" ID tubing. The pressure regulator is connected to the microfluidic device through 1/16" OD x 0.020" ID PFA tubing with 24 gauge metal tubing at its terminus.</li>
	<li>Rinse the fluidic connections and the microfluidic device with 0.5 mL of buffer solution to ensure that all the air bubbles are removed from the system including the outlet channels. Typical flow rates used for clearing bubbles range between 2000-5000 &mu;L/hr. After the air bubbles are rinsed out of the microfluidic channels, reduce the flow rate to 50-100 &mu;L/hr, which is a typical volumetric flow rate for particle trapping.</li>
	<li>At this point, the fluidic connections are established, the sample and the buffer solutions are delivered to the microfluidic device at a fixed flow rate (50-100 &mu;L/hr), and the device is ready for hydrodynamic trapping.</li>
</ol>
4. Hydrodynamic Trapping Procedure
<ol>
	<li>Run the custom-built LabVIEW code, which automates particle trapping (see Usage Note for LabVIEW code below).</li>
	<li>Using the microscope x-y translation stage, position the trapping region (cross-slot) at the center of the camera view. Bring the trapping region into focus of the objective lens and adjust the camera settings to optimize imaging conditions.</li>
	<li>Choose a rectangular region of interest (ROI) within the camera's field of view such that the center of the ROI will be the position of the trap center.</li>
	<li>Initialize the offset pressure applied to the on-chip valve. In one of the outlet channels, a 100-200 &mu;m wide constriction is introduced to provide an offset pressure for the on-chip valve. The constant off-set pressure enables the on-chip valve to adjust the stagnation point position in the vicinity of the center of the channel cross-slot. For most experiments, the offset pressure is set between 0-12 psi depending on the channel dimensions (height and width), the constriction width, and the specifications of the on-chip valve (valve size, membrane thickness, etc.).</li>
	<li>Initiate the feedback controller and adjust the proportional gain to optimize trap response. The feedback controller will adjust the pressure applied to the on-chip valve in order to move the stagnation point position, which minimizes the error or the distance between the particle position and the set point (trap center). Depending on the flow rate and the on-chip valve position, there is an optimal proportional gain value, which increases trap stability and eliminates unwanted particle oscillations.</li>
	<li>Trap a particle. The LabVIEW code will automatically trap one of the particles entering the trapping region. Once a desired particle is trapped, it is possible to shut off the sample flow and isolate the trapped particle in buffer solution, if desired.</li>
	<li>Monitor the trapped particle and maintain particle focus within the image plane using manual focus or an automated focus microscope setup. It may be necessary to slightly adjust the proportional gain of the feedback controller in order to ensure trap stability during the course of a long time-scale trapping event (minutes to hours).</li>
</ol>
LabVIEW Code: Usage Note for Feedback Controller
Automated particle trapping is achieved using a linear feedback control algorithm implemented using a custom LabVIEW code. The LabVIEW code captures images from a CCD camera and transmits an electric potential (voltage) to a pressure regulator, which actively modulates the position (partially open/closed state) of an on-chip dynamic pneumatic valve. As the valve position changes, the hydrodynamic flow rate in one outlet line is adjusted, thereby re-positioning the stagnation point and enabling hydrodynamic trapping. The steps in the feedback loop are sequentially and iteratively executed at the rate of image capturing (10-60 Hz). The LabVIEW code executes the following steps during each feedback loop cycle:
<ul>
	<li>Image capture. An image is acquired for a "target" particle in the trapping region of the microfluidic device using fluorescence microscopy with a 10x objective lens (NA: 0.4) and a CCD camera.</li>
	<li>Particle tracking. Particle centroid position is determined, and the particle tracking algorithm is initiated. Particles are localized by fitting the emission intensity profile of the particle to a point spread function (PSF), from which the centroid position is determined.</li>
	<li>Flow field control. The updated pressure intended for the on-chip dynamic valve is calculated using a feedback control algorithm with a proportional controller. In this way, the action of the valve is to re-position the stagnation point, which exerts a hydrodynamic force on the particle in order to steer the particle toward the trap center.</li>
</ul>
The LabVIEW code records the following data for every image captured during particle trapping: 1) time elapsed, 2) centroid (x,y) position of the trapped particle, 3) position of the trap center, 4) distance of the particle from the trap center, 5) pressure applied to the on-chip valve. In addition, the code also records a movie of the trapped particle in AVI file format.
5. Representative Results
We trapped fluorescent polystyrene beads of various size (100, 540, 830 nm, and 2.2 &mu;m diameter) using a hydrodynamic trap. Figure 1(a) shows an image of a particle trapped at the cross-slot junction in a microfluidic device. The trajectory of a trapped particle may be determined directly from the centroid position data recorded by the LabVIEW code during a trapping event or by tracking and localizing the trapped particle from the recorded movie file. Figure 1(b) shows the trajectory of a trapped particle (2.2 &mu;m fluorescent polystyrene bead) along the outlet channel direction. The bead is initially trapped (squares) for 3 min and is then released from the trap and escapes along one of the outlet channels (circles). Particle trajectories along the compressional flow axis (inlet channel direction; data not shown) are similar to particle trajectories along the extensional flow axis (outflow direction) as shown in Figure 1(b). A histogram of particle displacement from the trap center for a trapped bead (2.2 &mu;m diameter) along the outlet channel directions is shown in Figure 1(c). Using the feedback control algorithm described in this work, trapped particles are confined to within &plusmn;1 &mu;m of the trap center along the inlet and outlet channel directions.
A schematic of the microfluidic device used for hydrodynamic trapping is shown in Figure 2. The integrated microfluidic device consists of a fluidic layer and a control layer and is fabricated using standard multilayer soft lithography as described in this article. The fluidic layer contains the buffer and sample channels, as well as the cross-slot channel geometry to facilitate hydrodynamic trapping. The control layer consists of a pneumatic valve positioned above one of the outlet channels in the fluidic layer, and the control and fluidic layers are separated by a thin elastomeric membrane. During device operation, the valve in the control layer is pressurized with nitrogen gas, which forces the thin membrane into the fluidic layer, thereby inducing a constriction in the outlet channel. The dynamic pneumatic valve constricts the outlet channel by variable amounts by changing the pressure applied to the control layer, which adjusts the relative flow rates in the outlet channels and enables fine-scale control of the stagnation point.
<img src="/files/ftp_upload/2517/2517fig1.jpg" alt="Figure 1" /> Figure 1: Particle Trapping. (a) Image of a single bead confined in the hydrodynamic trap. In addition to the bead at the trap center, several untrapped beads are shown in the trapping region. (b) Trajectory of a trapped particle along the outlet channels (squares). When the particle is released from the trap (arrow), it escapes along one of the outlet channels (circles). (c) Histogram of displacements of a trapped bead (2.2 &mu;m diameter) from the trap center along the outlet channels.
<img src="/files/ftp_upload/2517/2517fig2.jpg" alt="Figure 2" /> Figure 2: Schematic of microfluidic device for hydrodynamic trapping. The hydrodynamic trap is constructed using a two-layer microfluidic device. The fluidic layer consists of a sample inlet, four buffer inlets, and two waste outlets. The control layer consists of a pneumatic membrane valve situated on top of one of the outlet channels in the fluidic layer. A constriction in the opposing outlet channel provides an offset pressure for the pneumatic valve. Typical channel dimensions range between 100-500 &mu;m. In region (A), sample inlet is flow focused by two buffer inlets. In region (B), opposing inlet streams converge at the cross-slot junction where trapping occurs. The pneumatic valve (C) is positioned on top of one of the outlet channels. The stagnation point position is modulated by regulating pressure to this valve.

<ol>
	<li>Tanyeri, M., Johnson-Chavarria, E. M., Schroeder, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrodynamic+Trap+for+Single+Particles+and+Cells.">Hydrodynamic Trap for Single Particles and Cells.</a> Applied Physics Letters. 96, 224101-224101 (2010).</li><li>Ashkin, A., Dziedzic, J. M., Bjorkholm, J. E., Chu, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Observation+of+a+Single-Beam+Gradient+Force+Optical+Trap+for+Dielectric+Particles.">Observation of a Single-Beam Gradient Force Optical Trap for Dielectric Particles.</a> Optics Letters. 11, 288-290 (1986).</li><li>Neuman, K. C., Block, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+trapping.">Optical trapping.</a> Review of Scientific Instruments. 75, 2787-2809 (2004).</li><li>Gosse, C., Croquette, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnetic+tweezers:+Micromanipulation+and+force+measurement+at+the+molecular+level.">Magnetic tweezers: Micromanipulation and force measurement at the molecular level.</a> Biophysical Journal. 82, 3314-3329 (2002).</li><li>Chiou, P. Y., Ohta, A. T., Wu, M. C., C, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Massively+parallel+manipulation+of+single+cells+and+microparticles+using+optical+images.">Massively parallel manipulation of single cells and microparticles using optical images.</a> Nature. 436, 370-372 (2005).</li><li>Cohen, A. E., Moerner, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+for+trapping+and+manipulating+nanoscale+objects+in+solution.">Method for trapping and manipulating nanoscale objects in solution.</a> Applied Physics Letters. 86, 093109-09 (2005).</li><li>Evander, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noninvasive+acoustic+cell+trapping+in+a+microfluidic+perfusion+system+for+online+bioassays&amp;quot;,+Analytical+Chemistry+79.">Noninvasive acoustic cell trapping in a microfluidic perfusion system for online bioassays&amp;quot;, Analytical Chemistry 79.</a> , 2984-2991 (2007).</li><li>Unger, M. A., Chou, H. P., Thorsen, T., Scherer, A., Quake, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monolithic+microfabricated+valves+and+pumps+by+multilayer+soft+lithography.">Monolithic microfabricated valves and pumps by multilayer soft lithography.</a> Science. 288, 113-116 (2000).</li><li>Kim, M. C., Wang, Z. H., Lam, R. H. W., Thorsen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Building+a+better+cell+trap:+Applying+Lagrangian+modeling+to+the+design+of+microfluidic+devices+for+cell+biology.">Building a better cell trap: Applying Lagrangian modeling to the design of microfluidic devices for cell biology.</a> Journal of Applied Physics. 103, (2008).</li><li>Lutz, B. R., Chen, J., Schwartz, D. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrodynamic+tweezers:+1.+Noncontact+trapping+of+single+cells+using+steady+streaming+microeddies.">Hydrodynamic tweezers: 1. Noncontact trapping of single cells using steady streaming microeddies.</a> Analytical Chemistry. 78, 5429-5435 (2006).</li></ol>

No conflicts of interest declared.

Current microfluidic methods for particle manipulation based on hydrodynamic flow can be characterized as contact-based or non-contact methods. Contact-based methods use fluid flow to physically confine and immobilize particles against microfabricated channel walls 9, whereas non-contact methods rely on circulating flow or microeddies 10. In this work, we present a method for free-solution particle trapping using the sole action of fluid flow. The hydrodynamic trap enables confinement and manipulati...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>21 gauge blunt needle</td>
<td>&nbsp;</td>
<td> Zephyrtronics</td>
<td> ZT-5-021-1-L</td>
<td>For punching port holes in PDMS</td>
</tr>
<tr>
<td>3 ml plastic syringe</td>
<td>&nbsp;</td>
<td>BD</td>
<td>309585</td>
<td>For filling valve with oil</td>
</tr>
<tr>
<td>Si wafers</td>
<td>&nbsp;</td>
<td>University Wafer</td>
<td>&nbsp;</td>
<td> 3&#x201D; P(100) single side polished 380 &mu;m test grade</td>
</tr>
<tr>
<td>Cover glass</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>48404-428</td>
<td>24 x 40 mm #1.5</td>
</tr>
<tr>
<td>DAQ card</td>
<td>&nbsp;</td>
<td>National Instruments</td>
<td>PCI 6229</td>
<td>&nbsp;</td>
<tr>
<td>Fluorescent beads</td>
<td>&nbsp;</td>
<td>Spherotech</td>
<td>FP-2056-2</td>
<td>2.2 &mu;m Nile red</td>
</tr>
<tr>
<td>Fluorinert </td>
<td>&nbsp;</td>
<td>3M</td>
<td>FC 40</td>
<td>Fluorinated carrier oil</td>
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<tr>
<td>Inverted Microscope</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>IX-71</td>
<td>&nbsp;</td>
<tr>
<td>LabVIEW</td>
<td>&nbsp;</td>
<td>National Instruments</td>
<td>Version 9.0f3 (32bit)</td>
<td>&nbsp;</td>
<tr>
<td>Stereo Microscope</td>
<td>&nbsp;</td>
<td>Leica</td>
<td>MZ6</td>
<td>For aligning PDMS control layer to fluidic layer.</td>
</tr>
<tr>
<td>Mechanical Convection Oven</td>
<td>&nbsp;</td>
<td> VWR</td>
<td>1300U</td>
<td>For baking devices to create monolithic PDMS slabs with two layers.</td>
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<tr>
<td>Microfluidic tubing and connectors</td>
<td>&nbsp;</td>
<td>Upchurch Scientific</td>
<td>&nbsp;</td>
<td>1/16 x .020 PFA tubing and super flangeless fittings</td>
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<td>&nbsp;</td>
<td>GE Silicones</td>
<td>RTV 615 A&amp;B</td>
<td>&nbsp;</td>
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<td>Plasma Chamber</td>
<td>&nbsp;</td>
<td>Harrick</td>
<td>PDC-001</td>
<td>&nbsp;</td>
<tr>
<td>Pressure Transducer</td>
<td>&nbsp;</td>
<td>Proportion Air</td>
<td>DQPV1</td>
<td>&nbsp;</td>
<tr>
<td>Spin Coater</td>
<td>&nbsp;</td>
<td>Specialty Coating Systems</td>
<td>G3P-8 Spin Coat</td>
<td>&nbsp;</td>
<tr>
<td>Photoresist</td>
<td>&nbsp;</td>
<td>MicroChem</td>
<td>SU 8 2050</td>
<td>&nbsp;</td>
<tr>
<td>Syringe Pump</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>PHD 2000 Programmable</td>
<td>&nbsp;</td>
<tr>
<td>Terminal Block</td>
<td>&nbsp;</td>
<td>National Instruments</td>
<td>BNC 2110</td>
<td>For analog output to pressure regulator and read out.</td>
</tr>
<tr>
<td>UV Collimated Light Source and Exposure System</td>
<td>&nbsp;</td>
<td>OAI</td>
<td>Model 30 Enhanced Light Source</td>
<td>&nbsp;</td>		</tbody>
		</table>
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a microfluidic-based hydrodynamic trap for single particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

Watch this Scientific Journal Video about A Microfluidic-based Hydrodynamic Trap for Single Particles at JoVE.com

A Microfluidic-based Hydrodynamic Trap for Single Particles

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

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16.6K Görüntüleme.  University of Illinois at Urbana-Champaign. Bu makalede, hidrodinamik akış dayalı parçacık hapsi mikroakışkan tabanlı bir yöntem mevcut. Biz böylece entegre bir microdevice keyfi parçacıkların hapsi ve mikromanipülasyon sağlayan bir geribildirim kontrol mekanizması kullanılarak bir sıvı durgunluk noktasında kararlı parçacık yakalama göstermektedir.

Video: A Microfluidic-based Hydrodynamic Trap for Single Particles

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and pathways as well as its ease of cultivation. Several C. elegans disease models have been reported, including neurodegenerative disorders such as Parkinson's disease (PD), which involves the degeneration of dopaminergic (DA) neurons 1. Both transgenes and neurotoxic chemicals have been used to induce DA neurodegeneration and consequent movement defects in worms, allowing for investigations into the basis of neurodegeneration and screens for neuroprotective genes and compounds 2,3.
Screens in lower eukaryotes like C. elegans provide an efficient and economical means to identify compounds and genes affecting neuronal signaling. Conventional screens are typically performed manually and scored by visual inspection; consequently, they are time-consuming and prone to human errors. Additionally, most focus on cellular level analysis while ignoring locomotion, which is an especially important parameter for movement disorders.
We have developed a novel microfluidic screening system (Figure 1) that controls and quantifies C. elegans' locomotion using electric field stimuli inside microchannels. We have shown that a Direct Current (DC) field can robustly induce on-demand locomotion towards the cathode ("electrotaxis") 4. Reversing the field's polarity causes the worm to quickly reverse its direction as well. We have also shown that defects in dopaminergic and other sensory neurons alter the swimming response 5. Therefore, abnormalities in neuronal signaling can be determined using locomotion as a read-out. The movement response can be accurately quantified using a range of parameters such as swimming speed, body bending frequency and reversal time.
Our work has revealed that the electrotactic response varies with age. Specifically, young adults respond to a lower range of electric fields and move faster compared to larvae 4. These findings led us to design a new microfluidic device to passively sort worms by age and phenotype 6.
We have also tested the response of worms to pulsed DC and Alternating Current (AC) electric fields. Pulsed DC fields of various duty cycles effectively generated electrotaxis in both C. elegans and its cousin C. briggsae 7. In another experiment, symmetrical AC fields with frequencies ranging from 1 Hz to 3 KHz immobilized worms inside the channel 8.
Implementation of the electric field in a microfluidic environment enables rapid and automated execution of the electrotaxis assay. This approach promises to facilitate high-throughput genetic and chemical screens for factors affecting neuronal function and viability.

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans&#039; Locomotion

A semi-automated micro-electro-fluidic method to induce on-demand locomotion in Caenorhabditis elegans is described. This method is based on the neurophysiologic phenomenon of worms responding to mild electric fields (&#x201C;electrotaxis&#x201D;) inside microfluidic channels. Microfluidic electrotaxis serves as a rapid, sensitive, low-cost, and scalable technique to screen for factors affecting neuronal health.

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and ...

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

A Microfluidic-based Electrochemical Biochip for Label-free DNA Hybridization Analysis

Miniaturization of analytical benchtop procedures into the micro-scale provides significant advantages in regards to reaction time, cost, and integration of pre-processing steps. Utilizing these devices towards the analysis of DNA hybridization events is important because it offers a technology for real time assessment of biomarkers at the point-of-care for various diseases. However, when the device footprint decreases the dominance of various physical phenomena increases. These phenomena influence the fabrication precision and operation reliability of the device. Therefore, there is a great need to accurately fabricate and operate these devices in a reproducible manner in order to improve the overall performance. Here, we describe the protocols and the methods used for the fabrication and the operation of a microfluidic-based electrochemical biochip for accurate analysis of DNA hybridization events. The biochip is composed of two parts: a microfluidic chip with three parallel micro-channels made of polydimethylsiloxane (PDMS), and a 3 x 3 arrayed electrochemical micro-chip. The DNA hybridization events are detected using electrochemical impedance spectroscopy (EIS) analysis. The EIS analysis enables monitoring variations of the properties of the electrochemical system that are dominant at these length scales. With the ability to monitor changes of both charge transfer and diffusional resistance with the biosensor, we demonstrate the selectivity to complementary ssDNA targets, a calculated detection limit of 3.8 nM, and a 13% cross-reactivity with other non-complementary ssDNA following 20 min&#160;of incubation. This methodology can improve the performance of miniaturized devices by elucidating on the behavior of diffusion at the micro-scale regime and by enabling the study of DNA hybridization events.

We present a microfluidic-based electrochemical biochip for DNA hybridization detection. Following ssDNA probe functionalization, the specificity, sensitivity, and detection limit are studied with complementary and non-complementary ssDNA targets. Results illustrate the influence of the DNA hybridization events on the electrochemical system, with a detection limit of 3.8 nM.

Miniaturization of analytical benchtop procedures into the micro-scale provides significant advantages in regards to reaction time, cost, and integration ...

Ordering Single Cells and Single Embryos in 3D Confinement: A New Device for High Content Screening

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening based on cells in such environments have therefore serious limitations: cell organelles show extreme phenotypes, cell morphologies and sizes are heterogeneous and/or specific cell organelles cannot be properly visualized. In addition, cells in vivo are located in a 3D environment; in this situation, cells show different phenotypes mainly because of their interaction with the surrounding extracellular matrix of the tissue. In order to standardize and generate order of single cells in a physiologically-relevant 3D environment for cell-based assays, we report here the microfabrication and applications of a device for in vitro 3D cell culture. This device consists of a 2D array of microcavities (typically 105 cavities/cm2), each filled with single cells or embryos. Cell position, shape, polarity and internal cell organization become then normalized showing a 3D architecture. We used replica molding to pattern an array of microcavities, &#8216;eggcups&#8217;, onto a thin polydimethylsiloxane (PDMS) layer adhered on a coverslip. Cavities were covered with fibronectin to facilitate adhesion. Cells were inserted by centrifugation. Filling percentage was optimized for each system allowing up to 80%. Cells and embryos viability was confirmed. We applied this methodology for the visualization of cellular organelles, such as nucleus and Golgi apparatus, and to study active processes, such as the closure of the cytokinetic ring during cell mitosis. This device allowed the identification of new features, such as periodic accumulations and inhomogeneities of myosin and actin during the cytokinetic ring closure and compacted phenotypes for Golgi and nucleus alignment. We characterized the method for mammalian cells, fission yeast, budding yeast, C. elegans with specific adaptation in each case. Finally, the characteristics of this device make it particularly interesting for drug screening assays and personalized medicine.

We report a device and a new method to study cells and embryos. Single cells are precisely ordered in microcavity arrays. Their 3D confinement is a step towards 3D environments encountered in physiological conditions and allows organelle orientation. By controlling cell shape, this setup minimizes variability reported in standard assays.

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening ...

Cultivation of Mammalian Cells Using a Single-use Pneumatic Bioreactor System

Recent advances in mammalian, insect, and stem cell cultivation and scale-up have created tremendous opportunities for new therapeutics and personalized medicine innovations. However, translating these advances into therapeutic applications will require in vitro systems that allow for robust, flexible, and cost effective bioreactor systems. There are several bioreactor systems currently utilized in research and commercial settings; however, many of these systems are not optimal for establishing, expanding, and monitoring the growth of different cell types. The culture parameters most challenging to control in these systems include, minimizing hydrodynamic shear, preventing nutrient gradient formation, establishing uniform culture medium aeration, preventing microbial contamination, and monitoring and adjusting culture conditions in real-time. Using a pneumatic single-use bioreactor system, we demonstrate the assembly and operation of this novel bioreactor for mammalian cells grown on micro-carriers. This bioreactor system eliminates many of the challenges associated with currently available systems by minimizing hydrodynamic shear and nutrient gradient formation, and allowing for uniform culture medium aeration. Moreover, the bioreactor&#8217;s software allows for remote real-time monitoring and adjusting of the bioreactor run parameters. This bioreactor system also has tremendous potential for scale-up of adherent and suspension mammalian cells for production of a variety therapeutic proteins, monoclonal antibodies, stem cells, biosimilars, and vaccines.

Using a pneumatic bioreactor, we demonstrate the assembly, operation, and performance of this single-use bioreactor system for the growth of mammalian cells.

Recent advances in mammalian, insect, and stem cell cultivation and scale-up have created tremendous opportunities for new therapeutics and personalized ...

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. However, to achieve robust sample manipulation it is necessary to address device integration with the macroscale environment. To realize repeatable, sensitive particle separation with microfluidic devices, this protocol presents a complete automated and integrated microfluidic platform that enables precise processing of 0.15&#8211;1.5 ml samples using microfluidic devices. Important aspects of this system include modular device layout and robust fixtures resulting in reliable and flexible world to chip connections, and fully-automated fluid handling which accomplishes closed-loop sample collection, system cleaning and priming steps to ensure repeatable operation. Different microfluidic devices can be used interchangeably with this architecture. Here we incorporate an acoustofluidic device, detail its characterization, performance optimization, and demonstrate its use for size-separation of biological samples. By using real-time feedback during separation experiments, sample collection is optimized to conserve and concentrate sample. Although requiring the integration of multiple pieces of equipment, advantages of this architecture include the ability to process unknown samples with no additional system optimization, ease of device replacement, and precise, robust sample processing.

This protocol describes a system architecture for performing automated small volume (0.15&#8211;1.5 ml) particle separations using a microfluidic device, and discusses methods to optimize acoustofluidic device performance and operation.

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. ...

A Label-free Technique for the Spatio-temporal Imaging of Single Cell Secretions

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning of such signaling can lead to many disorders. To understand cell-to-cell signaling, it is essential to study the spatial and temporal nature of the secreted molecules from the cell without disturbing the local environment. Various assays have been developed to study protein secretion, however, these methods are typically based on fluorescent probes which disrupt the relevant signaling pathways. To overcome this limitation, a label-free technique is required.
In this paper, we describe the fabrication and application of a label-free localized surface plasmon resonance imaging (LSPRi) technology capable of detecting protein secretions from a single cell. The plasmonic nanostructures are lithographically patterned onto a standard glass coverslip and can be excited using visible light on commercially available light microscopes. Only a small fraction of the coverslip is covered by the nanostructures and hence this technique is well suited for combining common techniques such as fluorescence and bright-field imaging.
A multidisciplinary approach is used in this protocol which incorporates sensor nanofabrication and subsequent biofunctionalization, binding kinetics characterization of ligand and analyte, the integration of the chip and live cells, and the analysis of the measured signal. As a whole, this technology enables a general label-free approach towards mapping cellular secretions and correlating them with the responses of nearby cells.

Inter-cellular communication is critical for controlling various physiological activities within and outside the cell. This paper describes a protocol for measuring the spatio-temporal nature of single cell secretions. To achieve this, a multidisciplinary approach is used which integrates label-free nanoplasmonic sensing with live cell imaging.

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to spatially fractionate the sample based on a biological property of interest. For the resultant spatial distribution to be used as the assay readout, microfluidic devices are often subjected to microscopic analysis requiring complex instrumentation with higher cost and reduced portability. To address this limitation, we have developed an integrated electronic sensing technology for multiplexed detection of particles at different locations on a microfluidic chip. Our technology, called Microfluidic CODES, combines Resistive Pulse Sensing with Code Division Multiple Access to compress 2D spatial information into a 1D electrical signal. In this paper, we present a practical demonstration of the Microfluidic CODES technology to detect and size cultured cancer cells distributed over multiple microfluidic channels. As validated by the high-speed microscopy, our technology can accurately analyze dense cell populations all electronically without the need for an external instrument. As such, the Microfluidic CODES can potentially enable low-cost integrated lab-on-a-chip devices that are well suited for the point-of-care testing of biological samples.

We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to ...