The authors would like to acknowledge Lloyd Ung for constructive input in early versions of this technique, Dr. Jeremy Agresti for pressure cap design tips, and Dr. Dongping Qi for his help in fabricating the pressure cap. We are grateful to the laboratories of M. Teitell and P. Gunaratne for providing a variety of cell samples for testing. We are grateful to the National Science Foundation (CAREER Award DBI-1254185), the UCLA Jonsson Comprehensive Cancer Center, and the UCLA Clinical and Translational Science Institute for supporting this work.

1. Microfluidic Device Design
NOTE: The device design has four basic functional regions: entry port, cell filter, constriction array, and exit port (Figure 1). The overall design can be applied to a wide array of cell types, with minor adjustments to dimensions. Provided here are a few basic design recommendations along with device parameters that are effective for a selection of both primary and immortalized cells.
<ol>
	<li>Select the width of constriction array channels (Figure 1B) to be approximately 30-50% of the average cell diameter; this constriction-to-cell size ratio results in significant cell deformation but minimal clogging. Given a cell type that has not previously been tested, it is prudent to design a range of constriction widths from ~30-50% the cell diameter to find the optimum width. As with many types of microfluidic devices, over 120 device master molds can be patterned on a single 4 inch silicon wafer, enabling an array of different device designs to be fabricated in a single fabrication process.</li>
	<li>Select the channel height to be at least 50% of the cell diameter; this ensures the cell is confined in 2&#160;dimensions. To prevent channel collapse during bonding, ensure that the channel aspect ratio is no less than 1:10 (height:width) throughout the device. For channels that must exceed this aspect ratio, add support posts to prevent collapse. Space support posts at a distance that is at least twice the cell diameter as to not impede cell flow.</li>
	<li>Include a filter at the entry ports to remove debris and disaggregate cell clusters (Figure 1B). Adjust the size and spacing of the filter posts so that the distance separating the posts is approximately equal to one cell diameter. 
	NOTE: The lower limit of feature size is set by the resolution at which the lithography mask is printed. Ensure that all features are within the resolution limit specified by the mask supplier.</li>
</ol>
2. Supplies and Preparation
NOTE: Before commencing any experiment, the following items must be prepared. A schematic of the entire setup is given in Figure 1.
<ol>
	<li>Fabricate the device master using standard techniques for lithographic micromachining14,15. Verify the height of the patterned features using a profilometer. 
	NOTE: While masters can be easily fabricated in a cleanroom facility, some labs have developed inexpensive lithography methods for use in a semi-clean environment16,17.</li>
	<li>Prepare the air source for pressure-driven flow, using a tank of compressed air and sequence of air regulators and fittings (Figure 1A).
	<ol>
		<li>Set up an air supply tank and manual regulator. 
		NOTE: A composition of 5% CO2 in air maintains a similar environment as a typical cell culture incubator, but other gas mixtures can also be used.</li>
		<li>Set up an electronically controlled pressure regulator in-line with the manual regulator. Use an electro-pneumatic converter to regulate and maintain a stable pressure drop across the channels, with pressure fluctuations of less than 2 kPa. Use the simple code written in LabVIEW (Supplementary Information) to input the desired pressure. 
		NOTE: The electro-pneumatic converter uses an internal feedback loop to adjust the valve outlet pressure to match the specified pressure across the microfluidic device.</li>
		<li>Set up a pressurized chamber to drive flow of the cell suspension. Assemble a cell suspension chamber out of a standard flow cytometer tube and a machined cap that creates a pressure-tight seal on the tube. 
		NOTE: The pressure chamber cap contains two orifices: an inlet that connects to the compressed air tank, and an outlet through which cells flow from the pressurized chamber into the device (Figure 2).</li>
	</ol>
	</li>
	<li>Set up an inverted microscope fitted with a camera that has an acquisition rate of at least 100 frames per sec&#160;to capture images of the cells flowing through the constriction array. Interface the camera with a video capture card and a computer.</li>
	<li>Prepare a stock surfactant solution of 10% w/w Pluronic F-127 in phosphate buffered saline (PBS) as adding a small amount of F127 to the cell media solution will help to minimize cell adhesion to the PDMS walls. To prepare the F127 solution, vortex and heat gently at 37 &#176;C&#160;to dissolve the F127. Prior to use, pass the solution through a 0.2 &#181;m syringe filter and store at room temperature. Prepare the surfactant solution in advance as vortexing and filtering will generate bubbles that will persist for more than 24 hr.</li>
</ol>
3. Microfluidic Device Fabrication
<ol>
	<li>Fabricate PDMS block with microfluidic channels.
	<ol>
		<li>Mix PDMS thoroughly at a ratio of 1:10 (w/w) curing agent to base.</li>
		<li>Pour the mixture over the device master (Step 2.1). Degas in a bell jar with applied vacuum until the entrapped bubbles disappear, or for about 10-20 min. 
		NOTE: After this time, there may still be bubbles at the air-PDMS interface, which is normal; these will typically dissipate during baking. It is most critical to ensure that there are no remaining bubbles adjacent to the channels.</li>
		<li>Bake the degassed device at 65 &#176;C&#160;for 4 hr. To prevent channels from collapsing, bake the device overnight to further crosslink the PDMS for a device with a higher elastic modulus that is more resistant to channel collapse. However, note that extended baking also embrittles the PDMS and may lead to cracking when punching holes (Step 3.3).</li>
	</ol>
	</li>
	<li>Remove microfluidic devices from the master mold. Cut individual microfluidic devices out of the PDMS block with a razor blade and remove the device from the master. Start by gently lifting off one corner of the device; slow and gentle peeling, as opposed to lifting the PDMS block straight up, reduces the stress on both the PDMS and the master.</li>
	<li>Punch holes in the PDMS device to create connection ports for access between tubing and microchannels. Create holes using a biopsy punch, and&#160;punch from the channel side through to the exterior side of the device. 
	NOTE: A biopsy punch with a 0.75 mm bore size creates holes that interface perfectly with polyetheretherketone (PEEK) tubing (outer diameter = 1/32&#8221; or 0.79 mm) or polyethylene tubing (PE-20, outer diameter = 0.043&#8221; or 1.09 mm). Use a ring light to help visualize holes in devices with shallow channels18. Be sure the biopsy punch is sharp; after repeated use the cutting edge dulls and the biopsy punch should be discarded and replaced.</li>
	<li>Rinse the PDMS device with isopropanol (HPLC grade) to remove dust and lodged chunks of PDMS. Make sure the punched holes are clear of debris by directing a steady stream of pure isopropanol from a squeeze bottle through the holes. Blow dry with filtered air. After cleaning, place PDMS blocks with channels face up in a clean Petri dish to maintain device cleanliness. If required, store devices in this form until bonding.</li>
	<li>Clean the glass substrate by rinsing with methanol (HPLC grade), blow dry with filtered air, and place on&#160;a 200 &#176;C&#160;hotplate&#160;for 5-10 min to ensure the glass is completely clean and dry before plasma treatment. 
	NOTE: The glass substrate forms the bottom of each channel. Typically a glass slide suffices, since a 10X&#160;or 20X&#160;objective is ideal for imaging multiple channels in parallel. For higher resolution imaging, bond the device to a coverslip (#1.5 thickness).</li>
	<li>Bond the PDMS device to the glass substrate.
	<ol>
		<li>Oxygen plasma treat the channel side of the PDMS block and 1&#160;side of a glass slide. 
		NOTE: With a hand-held discharge unit, use a treatment time of 1 min, but optimize the treatment time for each oxygen plasma apparatus.</li>
		<li>Place the channel side of the PDMS on top of the treated side of the glass immediately after treatment.</li>
		<li>Hold together with light pressure for ~10 sec to promote bonding. Bake the entire device at an elevated temperature (60-80&#160;&#176;C) for at least 15 min to improve bond strength.</li>
	</ol>
	</li>
</ol>
4. Deforming Cells through Constricted Channels
<ol>
	<li>Inspect the microfluidic device under a microscope. Ensure that the channels are not collapsed or broken and that both inlet and outlet holes connect directly into the device channels by using a low-power objective (10X). Designate defective devices by scoring the surface of the PDMS with a razor blade and do not use them for experiments.</li>
	<li>Prepare the cell culture samples to be assayed. Culture cells using standard conditions.
	<ol>
		<li>For example, culture HL-60 cells in RPMI-1640 medium supplemented with 1% penicillin-streptomycin solution and 10% fetal bovine serum in 5% CO2 incubator at 37 &#176;C.</li>
		<li>For adherent cells, harvest them by trypsinization and resuspend in fresh growth medium. Use a standard trypsinization protocol; for example, add ~0.5 ml&#160;trypsin to a flask with 25 cm2 culture area, and resuspend in ~30 ml&#160;fresh medium. Differentiate HL-60 cells into neutrophil-type cells by all-trans retinoic (ATRA) at a final concentration of 5 &#181;M per 1 x 105 cells/ml as previously described19,20.</li>
		<li>Centrifuge the cells at 300 x g for 3 min, carefully aspirate the supernatant and resuspend cells to the appropriate final density in fresh culture medium with 0.1% v/v F-127.</li>
	</ol>
	</li>
	<li>Count the cells using a Coulter counter, hemacytometer, or flow cytometer. Adjust the cell suspension to a density of 1 x 106 to 5 x 106 cells/ml. Once the suspension density is set, supplement with approximately 0.1% v/v F-127. 
	NOTE: The cell density and surfactant concentration may need to be modified depending on the particular cell systems; Table 1&#160;provides a record of previously used concentrations of cells and F-127 for different cell types.</li>
	<li>Place cell suspension in a flow cytometer tube and connect to the pressure cap. Before connecting the tubing to the microfluidic device, adjust the pressure to about 14-21 kPa and flush until the cell suspension emerges from the tip of the tubing. This will help to minimize air bubbles in the tubing and device.</li>
	<li>Insert the tip of the tubing containing the cell suspension into the device inlet. If the device is bonded to a coverslip, place the device on a flat, solid surface such as the microscope stage before connecting the tubing; the coverslip is fragile and may otherwise break.</li>
	<li>Insert a piece of tubing into the exit port and route it into an empty tube for waste collection such as an empty Falcon tube taped to the side of the microscope stage. For consistency in the overall fluidic resistance of the device between experiments, ensure that the inlet and outlet tubing is of the same diameter and approximately the same length for each experiment.</li>
	<li>Gently ramp up the pressure to about 28 kPa or until cells flow through the channels. Position the device so that multiple channels are in the field of view and are perpendicular to the bottom of the screen. If the camera is limited by data size, acquire multiple videos to generate a sufficient number of independent data points. Adjust the frame rate as necessary for the desired data output. To capture changes in cell shape, use high speed imaging at 300 frames per sec&#160;(fps); to measure transit times of cells, use frame rates of approximately 100 fps.</li>
</ol>
5. Data Analysis
<ol>
	<li>Open the file Master_script.m (Downloadable Supplemental File) and run the program.</li>
	<li>Select the first video to be analyzed from the Windows Explorer window that appears. Specify the frame rate for the selected video and press enter. 
	NOTE: A figure will appear that prompts the user to select a rectangular cropping window for the first video.</li>
	<li>Select a window that encompasses all the channels from left to right and intersects the channels just above the first bulb of the array of channels at the top and bottom (Figure 3).</li>
	<li>Select the constriction regions from the cropped image. Pay special attention to maintain a fixed pixel distance between the top and bottom window boundary for every video selected. 
	NOTE: The top window edge specifies the topmost constriction and bottom window edge specifies the bottommost constriction (Figure 4); intermediate constrictions are approximated from this user input. Next, the algorithm displays the segmentation of cells for the first 50 frames of each video (Figure 5).</li>
	<li>Monitor the top left image (Figure 5A) closely to determine if the algorithm is accurately locating cells; a binarized image is superimposed on the source video to demonstrate the location of identified cells. 
	NOTE: The different windows, (Figures 5B-5D) are for code diagnosis and demonstrate the video frames after specific image processing operations.</li>
	<li>Select the next video to be processed from the Windows Explorer window. 
	NOTE: The algorithm will repeat steps 5.2-5.5 for each video selected.</li>
	<li>Select cancel once all desired videos have been added via the Windows Explorer window to instruct the function that the video list is complete. 
	NOTE: The algorithm will perform the remaining operations without user intervention. This will take approximately 1-2 hr&#160;depending on the number of videos processed and computer performance. Upon completion, the algorithm will produce a histogram of the transit time data and save the data in the directory as a MATLAB .mat file and a Microsoft Excel .xls file.</li>
</ol>

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The authors have no conflicts of interest to disclose.

Here we provide a comprehensive experimental procedure for analyzing the deformation of cells transiting through constricted microfluidic channels using pressure-driven flow. A MATLAB script enables automated data processing (Supplemental Material); an updated version of the code is maintained (<a href="http://www.ibp.ucla.edu/research/rowat" target="_blank">www.ibp.ucla.edu/research/rowat</a>). More broadly, the techniques presented here can be adapted in many cell-based microfluidic assays, including the effect of cyto...

Changes in cell shape are critical in numerous biological contexts. For example, erythrocytes and leukocytes deform through capillaries that are smaller than their own diameter1. In metastasis, cancer cells must deform through narrow interstitial gaps as well as tortuous vasculature and lymphatic networks to seed at secondary sites2. To probe the physical behavior of individual cells, microfluidic devices present an ideal platform that can be customized to study a range of cell behaviors including their ability to migrate through narrow gaps3 and to passively deform through micron-scale constrictions3-9. Polydimethylsiloxane (PDMS) microfluidic devices are optically transparent, enabling cell deformations to be visualized using light microscopy and analyzed using basic image processing tools. Moreover, arrays of constrictions can be precisely defined, enabling analysis of multiple cells simultaneously with a throughput that exceeds many existing techniques10,11.
Here we present a detailed experimental protocol for probing cell deformability using the &#8216;Cell Deformer&#8217; PDMS microfluidic device. The device is designed so that cells passage through sequential constrictions; this geometry is common in physiological contexts, such as the pulmonary capillary bed12. To gauge cell deformability, transit time provides a convenient metric that is easily measured as the time required for an individual cell to transit through a single constriction4,6. To maintain a constant pressure drop across the constricted channels during cell transit, we use pressure-driven flow. Our protocol includes detailed instructions on device design and fabrication, device operation by pressure-driven flow, preparation and imaging of cells, as well as image processing to measure the time for cells to deform through a series of constrictions. We include both device designs and vision data processing code as supplemental files. As a representative sample of data, we show cell transit time through a series of constrictions as a function of the number of constrictions passaged. Analysis of the timescale for cells to transit though narrow constrictions of a microfluidic device can reveal differences in the deformability of a variety of cell types4,5,13. The device demonstrated here uniquely surveys cell transit through a series of micron-scale constrictions; this design emulates the tortuous path that cells experience in circulation and also enables probing additional physical characteristics of the cells such as relaxation time.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:71px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Pluronic F-127 Block Copolymer Surfactant&#160;</td>
			<td style="width:71px;">Fisher Scientific&#160;</td>
			<td style="width:119px;">8409400</td>
			<td style="width:167px;">Produced by BASF, also available through Sigma</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">PDMS base and crosslinker</td>
			<td style="width:71px;">Essex Brownell</td>
			<td style="width:119px;">DC-184-1.1</td>
			<td style="width:167px;">Product commonly named Sylgard 184 Elastomer</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Oxygen plasma discharge unit</td>
			<td style="width:71px;">Enercon</td>
			<td style="width:119px;">Dyne-A-Mite 3D Treater</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Biopsy Punch, Harris Uni-Core (0.75 mm)</td>
			<td style="width:71px;">Ted Pella, Inc.</td>
			<td style="width:119px;">15072</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Fingertight Ferrule, 1/32&#34;</td>
			<td style="width:71px;">Upchurch Scientific</td>
			<td style="width:119px;">UP-F-113</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Fingertight III Fitting, 10-32</td>
			<td style="width:71px;">Upchurch Scientific</td>
			<td style="width:119px;">UP-F-300X</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">polyetheretherketone (PEEK) tubing, outer diameter = 1/32&#34;or 0.79 mm</td>
			<td style="width:71px;">Valco</td>
			<td style="width:119px;">TPK.515-25M</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">polyethylene (PE-20) tubing, 0.043&#34; or 1.09 mm</td>
			<td style="width:71px;">Becton Dickinson</td>
			<td style="width:119px;">427406</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Pressure regulator</td>
			<td style="width:71px;">Airgas or Praxair</td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Polyurethane tubing, 5/32&#8221; OD</td>
			<td style="width:71px;">McMaster Carr</td>
			<td style="width:119px;">5648K284</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Push-to-connect fittings</td>
			<td style="width:71px;">McMaster Carr</td>
			<td style="width:119px;">5111K91</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Voltage to Pressure (E/P) Electropneumatic Converter</td>
			<td style="width:71px;">Omega</td>
			<td style="width:119px;">IP413-020</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="52">
			<td height="52" style="height:52px;width:216px;">16-bit,250 kS/S, 80 Analog Inputs Multifunction DAQ</td>
			<td style="width:71px;">National Instruments</td>
			<td style="width:119px;">NI PCI 6225-779295-01</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="52">
			<td height="52" style="height:52px;width:216px;">Analog Connector Block-Screw Terminal</td>
			<td style="width:71px;">National Instruments</td>
			<td style="width:119px;">SCB-68-776844-01</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="52">
			<td height="52" style="height:52px;width:216px;">LabView System Design Software</td>
			<td style="width:71px;">National Instruments</td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="52">
			<td height="52" style="height:52px;width:216px;">Matlab Software</td>
			<td style="width:71px;">The MathWorks, Inc.</td>
			<td style="width:119px;">Matlab R2012a</td>
			<td style="width:167px;">Code requires the Image Processing Toolbox</td>
		</tr>
		<tr height="53">
			<td height="53" style="height:53px;width:216px;">Shielded Cable</td>
			<td style="width:71px;">National Instruments</td>
			<td style="width:119px;">SHC68-68</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

a microfluidic technique to probe cell deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

To investigate the deformability of different cell types, human myeloid leukemia cells (HL-60), differentiated neutrophil cells, mouse lymphocyte cells, and human ovarian cancer cell lines (OVCAR8, HEYA8) are evaluated using the &#8216;Cell Deformer&#8217; microfluidic technique. Representative results for the transit time of HL-60 and neutrophil-type HL-60 cells show the timescale for a single cell to transit through a series of constrictions, as shown in Figure 6. Transit time is measured for a populat...

Watch this Scientific Journal Video about A Microfluidic Technique to Probe Cell Deformability at JoVE.com

A Microfluidic Technique to Probe Cell Deformability

We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

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Video: A Microfluidic Technique to Probe Cell Deformability

Systematic Analysis of In Vitro Cell Rolling Using a Multi-well Plate Microfluidic System

A major challenge for cell-based therapy is the inability to systemically target a large quantity of viable cells with high efficiency to tissues of interest following intravenous or intraarterial infusion. Consequently, increasing cell homing is currently studied as a strategy to improve cell therapy. Cell rolling on the vascular endothelium is an important step in the process of cell homing and can be probed in-vitro using a parallel plate flow chamber (PPFC). However, this is an extremely tedious, low throughput assay, with poorly controlled flow conditions. Instead, we used a multi-well plate microfluidic system that enables study of cellular rolling properties in a higher throughput under precisely controlled, physiologically relevant shear flow1,2. In this paper, we show how the rolling properties of HL-60 (human promyelocytic leukemia) cells on P- and E-selectin-coated surfaces as well as on cell monolayer-coated surfaces can be readily examined. To better simulate inflammatory conditions, the microfluidic channel surface was coated with endothelial cells (ECs), which were then activated with tumor necrosis factor-&#945; (TNF-&#945;), significantly increasing interactions with HL-60 cells under dynamic conditions. The enhanced throughput and integrated multi-parameter software analysis platform, that permits rapid analysis of parameters such as rolling velocities and rolling path, are important advantages for assessing cell rolling properties in-vitro. Allowing rapid and accurate analysis of engineering approaches designed to impact cell rolling and homing, this platform may help advance exogenous cell-based therapy.

Systematic Analysis of In Vitro Cell Rolling Using a Multi-well Plate Microfluidic System

This study used a multi-well plate microfluidic system, significantly increasing throughput of cell rolling studies under physiologically relevant shear flow. Given the importance of cell rolling in the multi-step cell homing cascade and the importance of cell homing following systemic delivery of exogenous populations of cells in patients, this system offers potential as a screening platform to improve cell-based therapy.

A major challenge for cell-based therapy is the inability to systemically target a large quantity of viable cells with high efficiency to tissues of ...

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This technology has shown potential in addressing previously challenging applications; including, delivery to primary immune cells, cell reprogramming, carbon nanotube, and quantum dot delivery. This vector-free microfluidic platform relies on mechanical disruption of the cell membrane to facilitate cytosolic delivery of the target material. Herein, we describe the detailed method of use for these microfluidic devices including, device assembly, cell preparation, and system operation. This delivery approach requires a brief optimization of device type and operating conditions for previously unreported applications. The provided instructions are generalizable to most cell types and delivery materials as this system does not require specialized buffers or chemical modification/conjugation steps. This work also provides recommendations on how to improve device performance and trouble-shoot potential issues related to clogging, low delivery efficiencies, and cell viability.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This protocol provides detailed steps on how to use the system for a broad range of applications.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This ...

Analyzing Mixing Inhomogeneity in a Microfluidic Device by Microscale Schlieren Technique

In this paper, we introduce the use of microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. The microscale schlieren system is constructed from a Hoffman modulation contrast microscope, which provides easy access to the rear focal plane of the objective lens, by removing the slit plate and replacing the modulator with a knife-edge. The working principle of microscale schlieren technique relies on detecting light deflection caused by variation of refractive index1-3. The deflected light either escapes or is obstructed by the knife-edge to produce a bright or a dark band, respectively. If the refractive index of the mixture varies linearly with its composition, the local change in light intensity in the image plane is proportional to the concentration gradient normal to the optical axis. The micro-schlieren image gives a two-dimensional projection of the disturbed light produced by three-dimensional inhomogeneity.
To accomplish quantitative analysis, we describe a calibration procedure that mixes two fluids in a T-microchannel. We carry out a numerical simulation to obtain the concentration gradient in the T-microchannel that correlates closely with the corresponding micro-schlieren image. By comparison, a relationship between the grayscale readouts of the micro-schlieren image and the concentration gradients presented in a microfluidic device is established. Using this relationship, we are able to analyze the mixing inhomogeneity from associate micro-schlieren image and demonstrate the capability of microscale schlieren technique with measurements in a microfluidic oscillator4. For optically transparent fluids, microscale schlieren technique is an attractive diagnostic tool to provide instantaneous full-field information that retains the three-dimensional features of the mixing process.

Herein, we describe a procedure that employs microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. Through calibration, distribution of concentration gradient can be derived from the micro-schlieren image.

In this paper, we introduce the use of microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. The microscale schlieren ...

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular behaviors on disease-relevant time scales. Previously, microcontact printing has been used to fabricate two-dimensional functional arterial mimics through patterning of extracellular matrix protein as guidance cues for tissue organization. Vascular muscular thin films utilized these mimics to assess functional contractility. However, the microcontact printing fabrication technique used typically incorporates hydrophobic PDMS substrates. As the tissue turns over the underlying extracellular matrix, new proteins must undergo a conformational change or denaturing in order to expose hydrophobic amino acid residues to the hydrophobic PDMS surfaces for attachment, resulting in altered matrix protein bioactivity, delamination, and death of the tissues.
Here, we present a microfluidic deposition technique for patterning of the crosslinker compound genipin. Genipin serves as an intermediary between patterned tissues and PDMS substrates, allowing cells to deposit newly-synthesized extracellular matrix protein onto a more hydrophilic surface and remain attached to the PDMS substrates. We also show that extracellular matrix proteins can be patterned directly onto deposited genipin, allowing dictation of engineered tissue structure. Tissues fabricated with this technique show high fidelity in both structural alignment and contractile function of vascular smooth muscle tissue in a vascular muscular thin film model. This technique can be extended using other cell types and provides the framework for future study of chronic tissue- and organ-level functionality.

We present a method for microfluidic deposition of patterned genipin and fibronectin on PDMS substrates, allowing extended viability of vascular smooth muscle cell-dense tissues. This tissue fabrication method is combined with previous vascular muscular thin film technology to measure vascular contractility over disease-relevant time courses.

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

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 ...

Rapid Subtractive Patterning of Live Cell Layers with a Microfluidic Probe

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one particular implementation of the MFP, the hierarchical hydrodynamic flow confinement (hHFC), multiple liquids are simultaneously brought in contact with a substrate. Local chemical action and liquid shaping using the hHFC, is exploited to create cell patterns by locally lysing and removing cells. By utilizing the scanning ability of the MFP, user-defined patterns of cell monolayers are created. This protocol enables rapid, real-time and spatially controlled cell patterning, which can allow selective cell-cell and cell-matrix interaction studies.

We present a protocol to perform subtractive patterning of live cell monolayers on a surface. This is achieved by local and selective lysis of adherent cells using a microfluidic probe (MFP). The cell lysate retrieved from local regions can be used for downstream analysis, enabling molecular profiling studies.

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...

Large-area Scanning Probe Nanolithography Facilitated by Automated Alignment and Its Application to Substrate Fabrication for Cell Culture Studies

Scanning probe microscopy has enabled the creation of a variety of methods for the constructive (&#39;additive&#39;) top-down fabrication of nanometer-scale features. Historically, a major drawback of scanning probe lithography has been the intrinsically low throughput of single probe systems. This has been tackled by the use of arrays of multiple probes to enable increased nanolithography throughput. In order to implement such parallelized nanolithography, the accurate alignment of probe arrays with the substrate surface is vital, so that all probes make contact with the surface simultaneously when lithographic patterning begins. This protocol describes the utilization of polymer pen lithography to produce nanometer-scale features over centimeter-sized areas, facilitated by the use of an algorithm for the rapid, accurate, and automated alignment of probe arrays. Here, nanolithography of thiols on gold substrates demonstrates the generation of features with high uniformity. These patterns are then functionalized with fibronectin for use in the context of surface-directed cell morphology studies.

Here we present a protocol for wide-area scanning probe nanolithography enabled by the iterative alignment of probe arrays, as well as the utilization of lithographic patterns for cell-surface interaction studies.

Scanning probe microscopy has enabled the creation of a variety of methods for the constructive (&#39;additive&#39;) top-down fabrication of ...

A Novel Tenorrhaphy Suture Technique with Tissue Engineered Collagen Graft to Repair Large Tendon Defects

Surgical management of large tendon defects with tendon grafts is challenging, as there are a finite number of sites where donors can be readily identified and used. Currently, this gap is filled with tendon auto-, allo-, xeno-, or artificial grafts, but clinical methods to secure them are not necessarily translatable to animals because of the scale. In order to evaluate new biomaterials or study a tendon graft made up of collagen type 1, we have developed a modified suture technique to help maintain the engineered tendon in alignment with the tendon ends. Mechanical properties of these grafts are inferior to the native tendon. To incorporate engineered tendon into clinically relevant models of loaded repair, a strategy was adopted to offload the tissue engineered tendon graft and allow for the maturation and integration of the engineered tendon in vivo until a mechanically sound neo-tendon was formed. We describe this technique using incorporation of the collagen type 1 tissue engineered tendon construct.

In this paper, we present an in vitro and in situ protocol to repair a tendon gap of up to 1.5 cm by filling it with engineered collagen graft. This was performed by developing a modified suture technique to take the mechanical load until the graft matures into the host tissue.