This work was supported by a grant from the National Health Research Institutes (03-A1 BNMP11-014).

Note: The photomask designs for our microfluidic device fabrication were drawn by using a computer aided design (CAD) software. The designs were then utilized to fabricate chrome photomasks using a commercial service. The PDMS devices were made using soft lithography techniques.11
1. Fabrication of Master Molds by Lithography
<ol>
	<li>Before the photolithography process12, use the 4-inch silicon wafers as a substrate and dehydrate the wafers in a conventional oven at 120 &#176;C for 10 min.</li>
	<li>Clean the dehydrated silicon wafers by using oxygen plasma treatment at 100 watts for 30 sec in a plasma cleaner.</li>
	<li>Preheat two hotplates at 65 &#176;C and 95 &#176;C, respectively, for the following baking process.</li>
	<li>Coat 5 g of negative photoresist (PR) on the cleaned silicon wafers by a spin coater; spin at 1,200 rpm (SU-8 50) for 30 sec to produce the microchannel layer.</li>
	<li>Place the PR coated wafer on a preheated hotplate at 65 &#176;C for 12 min and transfer it to another preheated hotplate at 95 &#176;C for 33 min (for 100 &#181;m thick patterns) to perform a soft bake process.</li>
	<li>After baking, place the PR coated silicon wafer on the holder of a semi-automated mask aligner and align it to a 25,400 dpi resolution transparency photomask.</li>
	<li>Expose the PR coated silicon wafer to UV light (365 nm) at a dose of 500 mJ/cm2 to create the PR pattern on the silicon wafer.</li>
	<li>Remove the wafer from the aligner and place it on a hotplate for post-baking at 95 &#176;C for 12 min.</li>
	<li>Soak the wafers in SU-8 developer (propylene glycol monomethyl ether acetate, PGMEA) solution to wash off uncrosslinked PR for 12 min and gently dry with nitrogen gas to expose the alignment marks.</li>
	<li>Again, coat 5 g of negative photoresist on the wafers by a spin coater; spin at 700 rpm (SU-8 100) for 30 sec and 1,200 rpm (SU-8 10) for 30 sec for 300 &#181;m thick pattern and 27 &#181;m thick pattern respectively to make the microwell layer.</li>
	<li>Place the PR coated wafer on a hotplate at 65 &#176;C for 4 min and at 95 &#176;C for 8 min (for 27 &#181;m deep capture-well layer); and at 65 &#176;C for 40 min and at 95 &#176;C for 110 min (for 300 &#181;m deep culture-well layer).</li>
	<li>After cooling, place the PR coated silicon wafer on the mask aligner equipped with UV light.</li>
	<li>Expose the PR coated silicon wafer to the UV light (365 nm) at a dose of 250 mJ/cm2 (for 27 &#181;m thick pattern) and 700 mJ/cm2 (for 300 &#181;m thick pattern).</li>
	<li>Bake the wafers at 95 &#176;C for 5 min (27 &#181;m thick pattern) and 30 min (300 &#181;m thick pattern), respectively.</li>
	<li>Wash off the uncrosslinked PR by the PGMEA for 6 min (27 &#181;m thick pattern) and 25 min (300 &#181;m thick pattern), respectively, and then dry them with nitrogen gas.</li>
	<li>Measure the height of pattern features on the wafer with a scanning laser profilometer by placing the wafer on the xy-stage of a scanning laser profilometer.
	<ol>
		<li>Adjust the focal plane to clearly show the pattern features on the wafer by using the &#34;camera view&#34; observation mode under a 20X objective lens.</li>
		<li>Switch the observation mode from &#34;camera view&#34; to &#34;laser view&#34;, and set the upper and lower positions of the features.</li>
		<li>Set the measurement mode, area, quality, and z-pitch up to transparent (top), 1 line (1,024 x 1), high-accuracy and 0.5 &#181;m, respectively, and then press the start bottom to begin the measurement.</li>
	</ol>
	</li>
</ol>
2. Preparation of PDMS Devices for Single-cell Isolation
<ol>
	<li>Before PDMS casting, silanize the master molds with trichlorosilane to create a hydrophobic surface, which makes it easier to peel off the PDMS replicas from the master molds.
	<ol>
		<li>Place the master molds and a weighting boat containing 200 &#181;l of trichlorosilane in a desiccator, and apply vacuum (-85 kPa) for 15 min.</li>
		<li>Stop the vacuum and then leave the master molds in the desiccator to silanize the master molds at room temperature for at least 1 hr.</li>
		<li>Remove the master molds from the desiccator and place each master mold in a 10 cm Petri dish.</li>
	</ol>
	</li>
	<li>Mix a total amount of 17.6 g PDMS polymer kit containing a base and a curing agent at a ratio of 10:1, and then pour the PDMS onto the master mold in the Petri dish.</li>
	<li>Place the Petri dish in a desiccator and apply vacuum (-85 kPa) for 1 hr to remove air bubbles in the PDMS.</li>
	<li>Remove the Petri dish from the desiccator and place it in a conventional oven at 65 &#176;C for 3 - 6 hr to cure the PDMS.</li>
	<li>Remove the cured PDMS replicas from the master molds and punch two holes as an inlet and an outlet at the two ends of the microchannel on the capture-well array PDMS replica by a puncher with inner-diameter of 0.75 mm for the fluidic channel (Figure 2A).</li>
	<li>Use tape to clean the surface of the PDMS replicas, and then place the PDMS replicas in a plasma cleaner for brief oxygen plasma treatment (100 watts for 14 sec).</li>
	<li>Remove the PDMS replicas from the oxygen plasma machine.</li>
	<li>Align a top (containing the capture microwells) and a bottom PDMS (containing the culture microwells) replica by hand under a stereomicroscope and bring them into contact.</li>
	<li>Place the aligned PDMS replicas in an oven at 65 &#176;C for 24 hr to achieve permanent bonding between the PDMS replicas to form the final device.</li>
	<li>Soak the PDMS device in a deionized (DI)-water filled container and place the container in a desiccator under vacuum (-85 kPa) for 15 min to remove air from the microchannel of the PDMS device.</li>
	<li>Place the DI water-filled PDMS device in a tissue culture hood and use UV light (wavelength of light: 254 nm) to sterilize of the device for 30 min.</li>
	<li>Replace the DI water in the PDMS device with a 5% bovine serum albumin (BSA) in 1x PBS solution and incubate at 37 &#176;C for 30 min to prevent cells from sticking to the PDMS surface. 
	Note: The BSA coating is critical to improve the transfer efficiency of isolated cells from capture-well to culture-well.</li>
	<li>Replace the 5% BSA solution in the PDMS device with sterilized 1x PBS solution.</li>
</ol>
3. Preparation of Single-cell Suspension
<ol>
	<li>Culture Neural stem cell KT98, and human carcinoma cell A549 and MDA-MB-435 we in Petri dish in conventional cell culture incubator (37 &#176;C, 5% CO2, and 95% humidity) to prepare cells for the PDMS device cell experiment.</li>
	<li>Remove and discard the spent culture medium (DMEM basal medium supplied with 10% fetal bovine serum and 1% antibiotics) from the culture dish with cells grown to 70 - 80% confluence.</li>
	<li>Gently wash the cells with sterilized PBS three times.</li>
	<li>Remove and discard the PBS, and add 2 ml of a recombinant enzyme mixture with proteolytic, collagenolytic, and DNase activities (please see the Materials List for the detailed information).</li>
	<li>Incubate the culture dish at room temperature for 5 min, and then tap the culture dish to facilitate cell detachment.</li>
	<li>Add 4 ml of sterilized PBS to disperse cells, and then transfer the cell suspension to a 15 ml conical tube.</li>
	<li>Centrifuge the tube at 300 x g for 3 min and remove the supernatant.</li>
	<li>Gently resuspend the cell pellet in 1 ml sterilized PBS and count the live cell number using the standard Trypan Blue exclusion method13. 
	Note: This resuspension step is critical for preparing well-dissociated single-cell suspension to improve the single-cell isolation efficiency.</li>
</ol>
4. Single-cell Isolation and Clonal Culture
<ol>
	<li>Load 50 &#181;l of cell suspension at a concentration of 2.2 - 2.5 x106 cells/ml into the microchannel of the PDMS device via the device&#39;s outlet hole with a handheld pipette. 
	Note: The cell suspension loading pipette needs to be stopped and held at the first stop position to avoid introducing air bubbles into the microchannel.</li>
	<li>Load another 50 &#181;l of cell suspension through the inlet hole to evenly fill up the whole microchannel with cells.</li>
	<li>Seal the outlet hole with a plug (a 3 mm long, 1 mm diameter cut nylon fishing line) to avoid flow induced by hydrostatic pressure from the droplets on inlet and outlet holes. 
	Note: The plugs were UV sterilized inside a tissue culture hood prior to use.</li>
	<li>Fill up a 1 ml sterile syringe with culture medium, eject air bubbles from it, and set it up on a syringe pump.</li>
	<li>Connect the medium loaded syringe to the inlet hole of the PDMS device via a 23 G blunt needle and a poly-tetrafluoroethene (PTFE) tubing.</li>
	<li>Remove the plug from outlet hole and allow a 2-min time interval to allow the cells to settle into the single-cell capture-wells by gravitational force.</li>
	<li>Wash away the uncaptured cells with 300 &#181;l of culture medium at a flow rate of 600 &#181;l/min driven by the syringe pump.</li>
	<li>Wait for 2 min to stabilize the device, and seal the inlet and outlet holes with plugs to form a closed culture system.</li>
	<li>Flip the device by hand to transfer the captured single-cells to the culture microwells.</li>
	<li>Place the PDMS device in a 100-mm tissue culture dish, and add 10 ml of sterilized PBS around the device to avoid culture medium evaporation from the microchannel.</li>
	<li>Move the culture dish to a conventional cell culture incubator (37 &#176;C, 5% CO2, and 95% humidity) for clonal culture of the single-cells.</li>
</ol>
5. Culture Medium Replenishment
<ol>
	<li>After 1 d of culture, replace the culture medium in the PDMS device with fresh medium to improve cell proliferation.</li>
	<li>Place two droplets of culture medium on top of the PDMS device near the inlet and outlet holes areas to avoid introducing air bubbles into the microchannel while performing the next step.</li>
	<li>Punch two holes from the top of the PDMS device near the two ends of the microchannel as an inlet and an outlet for the microchannel. 
	Note: Do not punch through the whole two layers of the PDMS device; instead, just punch through the top layer of the PDMS.</li>
	<li>Connect a 1 ml plastic syringe containing fresh medium to the inlet via a 23 G blunt needle and PTFE tubing.</li>
	<li>Flow 120 &#181;l fresh medium into the device for 5 min to replace the old medium.</li>
	<li>Insert two plugs to seal the inlet and outlet holes, and return the device to the cell culture incubator.</li>
	<li>Refresh the culture medium every 2 d during the period of culture by removing the sealing plugs, followed by repeating steps 5.4 to 5.5.</li>
</ol>

<ol>
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Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+a+prostate+from+a+single+adult+stem+cell.">Generation of a prostate from a single adult stem cell.</a> Nature. 456 (7223), 804-808 (2008).</li><li>Shapiro, E., Biezuner, T., Linnarsson, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+sequencing-based+technologies+will+revolutionize+whole-organism+science.">Single-cell sequencing-based technologies will revolutionize whole-organism science.</a> Nat Rev Genet. 14 (9), 618-630 (2013).</li><li>Rettig, J. 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The authors declare that they have no competing financial interests.

Microwell-based device systems6,14 have been utilized for single-cell manipulation and analysis, such as large-scale single cell trapping6 and single hematopoietic stem cell proliferation15. Although well size, number, and shape can be adjusted for specific applications, the single-cell isolation efficiency is always compromised when the size of the well is increased.9,15
To overcome this limitation, Park et al. reported a microfluidic chip with triangular microwells that have a high single-cell trapping rate (58.34%), while the microwell size is enlarged to allow for cell spreading and growth. However, the microwells (with the length of a side ~50 &#181;m) could only support cell proliferation for up to 2 d.16 Our dual-well strategy allows single-cell loading in large microwells to not be limited by the probability of the Poisson distribution encountered in limiting dilution methods and open well systems3, as demonstrated by providing high single-cell isolation efficiency (more than 75%) in large microwells whose size was sufficient for cell proliferation for at least 7 d (Figure 4A). Due to its ability to highly efficiently perform single-cell isolation and clonal culture with a simple device and operation procedure, we envision that our microfluidic platform could constitute a useful tool for a broad range of applications, including cancer stem cell selection by clonogenic assay17 and high-throughput cardiotoxicity screening of drugs18,19.
A major limitation of our single-cell isolation and culture platform is that it does not form individually closed culture-wells to prevent crosstalk between cells cultured in the culture-wells. Therefore, the behavior of the cells may be affected by the paracrine secretion of other isolated single-cells in the microfluidic device. The other limitation of our platform is the need to prepare relatively high-density cell suspension for high efficiency single-cell isolation. This high cell consumption can limit the applications of rare cell isolation and culture, such as aqueous humor cells.
Before the device operation, it is important to inject DI water from the inlet hole to fill the microchannel and carefully observe any fluid leakage from the bonded PDMS device. Fluid leakage affects the flow condition in the microchannel thus could adversely affect the cell capture and transferring outcome. In addition, although our single-cell loading efficiency study results show very low cell loss after transferring the isolated cells from capture-wells to culture-wells, indicating that most cells could be effectively transferred by flipping the device, the transferring efficiency is decreased if the BSA blocking step (2.12) is not properly performed. Users may modify the BSA blocking protocol or use alternative surface modification methods for their specific cell experiment needs (e.g., increasing BSA concentration or coating time when a more adherent cell type is used). The dimensions of the cell capture-wells are the dominant factors, which need to be optimized for the best single-cell loading efficiency in capture-wells, depending on the size of the specific cell type of interest. High multiple-cells-in-a-capture-well events are due to having the capture well size too large for the cell type of interest, whereas low cell loading efficiency in the capture well is an indication of the size of the capture well being too small. It is also important to minimize cell clusters in the prepared single cell suspension, as cell clusters can decrease the efficiency of single-cell loading in the capture-wells.

Currently placing single cells individually in a culture space is commonly achieved by using limiting dilution or fluorescence-activated cell sorting (FACS). For many laboratories, limiting dilution is a convenient method, as it only requires a pipette and tissue culture plates, which are readily available. In this case, a cell suspension is serially diluted to an appropriate cell density, and then placed into culture wells by using a manual pipette. These compartmented single cells are then used for cell analysis, such as genetic heterogeneity screening1 and colony formation2. However, this method is low-throughput and labor-intensive, without utilizing a robotic arm for assistance, because the Poisson distribution nature of the limiting dilution method restricts single-cell events to a maximum probability of 37%3. FACS machines with an integrated robotic arm can overcome the limitation of Poisson distribution by accurately placing one single-cell in a culture well at a time4. However, the high mechanical shear stress (thus, lowered cell viability)5 and machine purchase and operational costs have limited its usage in many laboratories.
To overcome the above limitations, microscale devices have been developed to highly efficiently load single cells into microwells6. However, the microwells do not provide adequate space for the loaded cells to proliferate, due to the need of making the size of each microwell close to that of a single cell to maximize the single-cell loading probability. As culture assays are required in many cell-based applications (e.g., clonogenic assay7), larger microwells (from 90 - 650 &#181;m in diameter or in side length) have also been utilized to allow for extended cell cultures. However, like the limiting dilution method, they also possess low single cell loading efficiencies, ranging from 10 - 30%.8,9
Previously, we have developed a high-throughput microfluidic platform to isolate single cells in individual microwells and demonstrate its application in clonogenic assay of the isolated cells.10 The device was made with poly-dimethylsiloxane (PDMS), and comprises two sets of microwell arrays with different microwell sizes, which can largely improve the efficiency in loading a single cell in a microwell whose size is significantly larger than the cell. Notably, this &#34;dual-well&#34; concept allows the size of the culture area to be flexibly adjusted without affecting the single-cell capture efficiency, making it straightforward to adjust the design of the device to suit different cell types and applications. This high-efficiency method should be useful for long-term cell culture experiments for cell heterogeneity studies and monoclonal cell line establishment.

<table border="0" cellpadding="0" cellspacing="0" width="1323">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:401px;">AutoCAD software</td>
			<td style="width:225px;">Autodesk</td>
			<td style="width:235px;">AutoCAD LT 2011</td>
			<td style="width:463px;">Part No. 057C1-74A111-1001</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Silicon wafer&#160;</td>
			<td>Eltech corperation</td>
			<td style="width:235px;">SPE0039</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Conventional oven</td>
			<td>YEONG-SHIN company</td>
			<td style="width:235px;">ovp45</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plasma cleaner</td>
			<td>Nordson</td>
			<td style="width:235px;">AP-300</td>
			<td>Bench-Top Plasma Treatment System</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">SU-8 50 negative photoresist</td>
			<td>MicroChem</td>
			<td>Y131269</td>
			<td style="width:463px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">SU-8 100 negative photoresist</td>
			<td>MicroChem</td>
			<td>Y131273</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Spin coater</td>
			<td>Synrex Co., Ltd.</td>
			<td>SC-HMI 2&#34; ~ 6&#34;</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Hotplate</td>
			<td>YOTEC company</td>
			<td>YS-300S</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Msak aligner</td>
			<td>Deya Optronic CO.</td>
			<td>A1K-5-MDA</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">SU-8 developer</td>
			<td>Grand Chemical Companies</td>
			<td>GP5002-000000-72GC</td>
			<td>Propylene glycol monomethyl ether acetate</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scanning laser profilometer</td>
			<td>KEYENCE</td>
			<td>VK-X 100</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Trichlorosilane</td>
			<td>Gelest, Inc</td>
			<td>SIT8174.0</td>
			<td style="width:463px;">TRIDECAFLUORO-1,1,2,2-TETRAHYDROOCTYL.&#160; Hazardous. Corrosive to the respiratory tract., reacts violently with water.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Desiccator</td>
			<td>Bel-Art Products&#160;</td>
			<td>F42020-0000</td>
			<td>SPACE SAVER VACUUM DESICCATOR 190MM WHITE BASE</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polydimethylsiloxane (PDMS) kit</td>
			<td>Dow corning</td>
			<td>Sylgard 184</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Harris Uni-Core puncher</td>
			<td>Ted Pella Inc.</td>
			<td>15072</td>
			<td>with 0.75 mm inner-diameter</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Removable tape</td>
			<td>3M Company</td>
			<td>Scotch Removable Tape 811</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Stereomicroscope</td>
			<td>Leica Microsystems</td>
			<td>Leica E24</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bovine serum albumin (BSA)</td>
			<td>Bersing Technology</td>
			<td>ALB001.500</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMEM basal medium</td>
			<td>Gibco</td>
			<td style="width:235px;">12800-017</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fetal bovine serum</td>
			<td>Thermo Hyclone</td>
			<td>SH30071.03HI</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Antibiotics</td>
			<td>Biowest</td>
			<td>L0014-100</td>
			<td>Glutamine-Penicillin-Streptomycin</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Recombinant enzyme mixture</td>
			<td>Innovative cell technology</td>
			<td style="width:235px;">AM-105</td>
			<td>Accumax</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DiIC12(3) cell membrane dye</td>
			<td>BD Biosciences</td>
			<td style="width:235px;">354218</td>
			<td>Used as a cell tracker</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Syringe pump</td>
			<td>Harvard Apparatus</td>
			<td style="width:235px;">703007</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Plastic syringe (1 mL)</td>
			<td>BD Biosciences</td>
			<td style="width:235px;">309659</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:401px;">23 gauge blunt needles</td>
			<td>Ever Sharp Technology, Inc.</td>
			<td style="width:235px;">TD21</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Poly-tetrafluoroethene (PTFE) tubing</td>
			<td>Ever Sharp Technology, Inc.</td>
			<td style="width:235px;">TFT-23T</td>
			<td>&#160;inner diameter, 0.51 mm; outer diameter, 0.82 mm</td>
		</tr>
	</tbody>
</table>

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.

The microfluidic platform for single-cell isolation and culture comprises a microchannel (200 &#181;m in height) with two sets of microwell arrays (Figure 2A). The two sets of microwell arrays are termed as capture-well (25 &#181;m in diameter and 27 &#181;m in depth) and culture-well (285 &#181;m in diameter and 300 &#181;m in depth) for single-cell isolation and culture, respectively, and each capture-well is positioned at the center of a culture-well when seen from the top-view (Figure 2B). For device operation (the schematic operation flow is illustrated in Figure 1), the required equipment (syringe pump, tissue culture incubator, and microscopes) and supplies (syringe, pipette, and tubing) are commonly available in biological laboratories. Moreover, the portability and transparency of the platform allowed the cells to be easily observed and analyzed in a device with a conventional microscope during the cell culture experiment.
The capture efficiency of cells in the capture-wells after the washing step ranges from 67.80 &#177; 11.38% to 85.16 &#177; 1.91% (depending on cell type) (Figure 3A). Moreover, most of the capture-wells contain only a single cell for all three cell types (89.89% - 92.98%), which was confirmed by analyzing the number of loaded cells in the culture-wells (Figure 3B). The final single-cell isolation efficiency in the culture-wells was more than 76% for KT98 and MDA-MB-435 cells, but only 61% for A549 cells due to cell size differences among the tested cell types.
For cancer research, clonogenic assay of single-cells can be utilized to test drug resistance and proliferation rates of individual cell colonies. The cell culture and clonogenic assay of the isolated single-cells in our platform were conducted by refreshing the culture medium every 2 d. This demonstration highlights the applicability of this device for studying cellular heterogeneity at the single cell level by measuring the differences in cell survival and proliferation rates of individual cells.
The cellular heterogeneity and proliferation rate of the isolated cells were analyzed from daily-acquired images. The results showed that the isolated single cells could be cultured for 7 d and exhibited different growth patterns. For example, 13% of the isolated cells had no cell division and remained as one single cell (Figure 4B), while 17.5% of the cells divided into more than 15 cells and formed cell colonies (Figure 4A) during the culture. Cellular heterogeneity among the proliferating cells was also evidenced by their different growth patterns of forming two cells (4.3%), three cells (2.5%), and 4 - 14 cells (15%) in the cell culture experiment (Figure 4C).
These results indicated that the platform can be used for long-term cell culture, clonogenic assays, and cellular heterogeneity studies. Having a large number of individual cell colonies grown in a small footprint area also makes microscopic analysis of the cells easier.
<img alt="Figure 1" src="/files/ftp_upload/54105/54105fig1.jpg" /> 
Figure 1. Schematic illustration of operation flow for high-throughput single-cell isolation and culture in the microfluidic platform. The operation procedure for single-cell isolation and clonal culture includes 4 steps: i) load cells into the microchannel by a plastic tip and manual pipette; ii) wash away the uncaptured cells with fresh culture medium driven by a syringe pump; iii) after sealing the inlet and outlet openings with plugs, the isolated cells were transferred from capture-wells to culture-wells by flipping the device; and iv) perform clonal culture (clonogenic assay) in culture-wells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54105/54105fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54105/54105fig2.jpg" /> 
Figure 2. Photograph and SEM images of the microfluidic single-cell isolation and culture platform. (A) Microfluidic platform loaded with red dye that shows the microchannel and microwell structures for single-cell culture. (B) A representative image taken from a cut device, showing the side-view of three single-cell isolation and clonal culture units in the microfluidic platform. Scale bar: 100 &#181;m. SEM images of single-cell culture (C) and capture (D) structures (microwells in different sizes). The sizes of the microwells can be adjusted for specific cell types and culture strategies. Scale bar: 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54105/54105fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54105/54105fig3.jpg" /> 
Figure 3. KT98, A549, and MDA-MB-435 single-cell isolation efficiency in the microfluidic platform. (A) Efficiency of KT98, A549, and MDA-MB-435 cells captured in capture-wells after washing away the uncaptured cells. The capture efficiency of cells was from 67.80% to 85.16% in the three cell types. (B) The single-cell ratio in total culture-wells (slash bar) was more than 76% for KT98 and MDA-MB-435, but 61.63% for A549. (C) A representative image of individual KT98 single-cells (red, stained with cell tracker) isolated in large culture-wells for clonal culture. Scale bar: 300 &#181;m. (D) Cell number in cell occupied culture-wells of the KT98 cell model. The ratio of culture-wells was 77.3% with a single-cell, 16.31% with no cells, 5.96% with two cells, and 0.43% with more than three cells. Error bars are represented as &#177;SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54105/54105fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54105/54105fig4.jpg" /> 
Figure 4. Cell culture, clonogenic assay, and cellular heterogeneity study of isolated A549 cells in the microfluidic platform for 7 d. (A) An isolated single-cell grew and proliferated to a cell colony in culture-well. (B) An isolated single-cell survived, but did not divide, after 7 d of culture. Scale bar: 100 &#181;m. (C) The isolated single-cells exhibited heterogeneous growth patterns after being cultured for 7 d. Error bars are represented as &#177;SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54105/54105fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Microfluidic Platform for High-throughput Single-cell Isolation and Culture at JoVE.com

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

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 ...

a-microfluidic-platform-for-high-throughput-single-cell-isolation

Nanyang Technological University

Research

JoVE Journal

Bioengineering

11.2K Views.  National Health Research Institutes, Taiwan. 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. 

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

High-throughput Protein Expression Generator Using a Microfluidic Platform

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and high-fidelity measurements of large systems. Microfluidics promises to fulfill many of these requirements, such as performing high-throughput screening experiments on-chip, encompassing biochemical, biophysical, and cell-based assays1. Since the early days of microfluidics devices, this field has drastically evolved, leading to the development of microfluidic large-scale integration2,3. This technology allows for the integration of thousands of micromechanical valves on a single device with a postage-sized footprint (Figure 1). We have developed a high-throughput microfluidic platform for generating in vitro expression of protein arrays (Figure 2) named PING (Protein Interaction Network Generator). These arrays can serve as a template for many experiments such as protein-protein 4, protein-RNA5 or protein-DNA6 interactions.
The device consist of thousands of reaction chambers, which are individually programmed using a microarrayer. Aligning of these printed microarrays to microfluidics devices programs each chamber with a single spot eliminating potential contamination or cross-reactivity Moreover, generating microarrays using standard microarray spotting techniques is also very modular, allowing for the arraying of proteins7, DNA8, small molecules, and even colloidal suspensions. The potential impact of microfluidics on biological sciences is significant. A number of microfluidics based assays have already provided novel insights into the structure and function of biological systems, and the field of microfluidics will continue to impact biology.

We present a microfluidic approach for the expression of protein arrays. The device consists of thousands of reaction chambers controlled by micro-mechanical valves. The microfluidic device is mated to a microarray-printed gene library. These genes are then transcribed and translated on-chip, resulting in a protein array ready for experimental use.

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and ...

High Throughput Single-cell and Multiple-cell Micro-encapsulation

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement from a bulk fluid environment with applications in high throughput screening, cytometry, and mass spectrometry. We describe a method to not only encapsulate single cells, but to repeatedly capture a set number of cells (here we demonstrate one- and two-cell encapsulation) to study both isolation and the interactions between cells in groups of controlled sizes. By combining drop generation techniques with cell and particle ordering, we demonstrate controlled encapsulation of cell-sized particles for efficient, continuous encapsulation. Using an aqueous particle suspension and immiscible fluorocarbon oil, we generate aqueous drops in oil with a flow focusing nozzle. The aqueous flow rate is sufficiently high to create ordering of particles which reach the nozzle at integer multiple frequencies of the drop generation frequency, encapsulating a controlled number of cells in each drop. For representative results, 9.9 &mu;m polystyrene particles are used as cell surrogates. This study shows a single-particle encapsulation efficiency Pk=1 of 83.7% and a double-particle encapsulation efficiency Pk=2 of 79.5% as compared to their respective Poisson efficiencies of 39.3% and 33.3%, respectively. The effect of consistent cell and particle concentration is demonstrated to be of major importance for efficient encapsulation, and dripping to jetting transitions are also addressed. 
Introduction 
Continuous media aqueous cell suspensions share a common fluid environment which allows cells to interact in parallel and also homogenizes the effects of specific cells in measurements from the media. High-throughput encapsulation of cells into picoliter-scale drops confines the samples to protect drops from cross-contamination, enable a measure of cellular diversity within samples, prevent dilution of reagents and expressed biomarkers, and amplify signals from bioreactor products. Drops also provide the ability to re-merge drops into larger aqueous samples or with other drops for intercellular signaling studies.1,2 The reduction in dilution implies stronger detection signals for higher accuracy measurements as well as the ability to reduce potentially costly sample and reagent volumes.3 Encapsulation of cells in drops has been utilized to improve detection of protein expression,4 antibodies,5,6 enzymes,7 and metabolic activity8 for high throughput screening, and could be used to improve high throughput cytometry.9 Additional studies present applications in bio-electrospraying of cell containing drops for mass spectrometry10 and targeted surface cell coatings.11 Some applications, however, have been limited by the lack of ability to control the number of cells encapsulated in drops. Here we present a method of ordered encapsulation12 which increases the demonstrated encapsulation efficiencies for one and two cells and may be extrapolated for encapsulation of a larger number of cells.
To achieve monodisperse drop generation, microfluidic "flow focusing" enables the creation of controllable-size drops of one fluid (an aqueous cell mixture) within another (a continuous oil phase) by using a nozzle at which the streams converge.13 For a given nozzle geometry, the drop generation frequency f and drop size can be altered by adjusting oil and aqueous flow rates Qoil and Qaq. As the flow rates increase, the flows may transition from drop generation to unstable jetting of aqueous fluid from the nozzle.14 
When the aqueous solution contains suspended particles, particles become encapsulated and isolated from one another at the nozzle. For drop generation using a randomly distributed aqueous cell suspension, the average fraction of drops Dk containing k cells is dictated by Poisson statistics, where Dk = &lambda;k exp(-&lambda;)/(k!) and &lambda; is the average number of cells per drop. The fraction of cells which end up in the "correctly" encapsulated drops is calculated using Pk = (k x Dk)/&Sigma;(k' x Dk'). The subtle difference between the two metrics is that Dk relates to the utilization of aqueous fluid and the amount of drop sorting that must be completed following encapsulation, and Pk relates to the utilization of the cell sample. As an example, one could use a dilute cell suspension (low &lambda;) to encapsulate drops where most drops containing cells would contain just one cell. While the efficiency metric Pk would be high, the majority of drops would be empty (low Dk), thus requiring a sorting mechanism to remove empty drops, also reducing throughput.15
Combining drop generation with inertial ordering provides the ability to encapsulate drops with more predictable numbers of cells per drop and higher throughputs than random encapsulation. Inertial focusing was first discovered by Segre and Silberberg16 and refers to the tendency of finite-sized particles to migrate to lateral equilibrium positions in channel flow. Inertial ordering refers to the tendency of the particles and cells to passively organize into equally spaced, staggered, constant velocity trains. Both focusing and ordering require sufficiently high flow rates (high Reynolds number) and particle sizes (high Particle Reynolds number).17,18 Here, the Reynolds number Re =uDh/&nu; and particle Reynolds number Rep =Re(a/Dh)2, where u is a characteristic flow velocity, Dh [=2wh/(w+h)] is the hydraulic diameter, &nu; is the kinematic viscosity, a is the particle diameter, w is the channel width, and h is the channel height. Empirically, the length required to achieve fully ordered trains decreases as Re and Rep increase. Note that the high Re and Rep requirements (for this study on the order of 5 and 0.5, respectively) may conflict with the need to keep aqueous flow rates low to avoid jetting at the drop generation nozzle. Additionally, high flow rates lead to higher shear stresses on cells, which are not addressed in this protocol. The previous ordered encapsulation study demonstrated that over 90% of singly encapsulated HL60 cells under similar flow conditions to those in this study maintained cell membrane integrity.12 However, the effect of the magnitude and time scales of shear stresses will need to be carefully considered when extrapolating to different cell types and flow parameters. The overlapping of the cell ordering, drop generation, and cell viability aqueous flow rate constraints provides an ideal operational regime for controlled encapsulation of single and multiple cells.
Because very few studies address inter-particle train spacing,19,20 determining the spacing is most easily done empirically and will depend on channel geometry, flow rate, particle size, and particle concentration. Nonetheless, the equal lateral spacing between trains implies that cells arrive at predictable, consistent time intervals. When drop generation occurs at the same rate at which ordered cells arrive at the nozzle, the cells become encapsulated within the drop in a controlled manner. This technique has been utilized to encapsulate single cells with throughputs on the order of 15 kHz,12 a significant improvement over previous studies reporting encapsulation rates on the order of 60-160 Hz.4,15 In the controlled encapsulation work, over 80% of drops contained one and only one cell, a significant efficiency improvement over Poisson (random) statistics, which predicts less than 40% efficiency on average.12
In previous controlled encapsulation work,12 the average number of particles per drop &lambda; was tuned to provide single-cell encapsulation. We hypothesize that through tuning of flow rates, we can efficiently encapsulate any number of cells per drop when &lambda; is equal or close to the number of desired cells per drop. While single-cell encapsulation is valuable in determining individual cell responses from stimuli, multiple-cell encapsulation provides information relating to the interaction of controlled numbers and types of cells. Here we present a protocol, representative results using polystyrene microspheres, and discussion for controlled encapsulation of multiple cells using a passive inertial ordering channel and drop generation nozzle.

Combining monodisperse drop generation with inertial ordering of cells and particles, we describe a method to encapsulate a desired number of cells or particles in a single drop at kHz rates. We demonstrate efficiencies twice exceeding those of unordered encapsulation for single- and double-particle drops.

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement ...

CometChip: A High-throughput 96-Well Platform for Measuring DNA Damage in Microarrayed Human Cells

DNA damaging agents can promote aging, disease and cancer and they are ubiquitous in the environment and produced within human cells as normal cellular metabolites. Ironically, at high doses DNA damaging agents are also used to treat cancer. The ability to quantify DNA damage responses is thus critical in the public health, pharmaceutical and clinical domains. Here, we describe a novel platform that exploits microfabrication techniques to pattern cells in a fixed microarray. The &#8216;CometChip&#8217; is based upon the well-established single cell gel electrophoresis assay (a.k.a. the comet assay), which estimates the level of DNA damage by evaluating the extent of DNA migration through a matrix in an electrical field. The type of damage measured by this assay includes abasic sites, crosslinks, and strand breaks. Instead of being randomly dispersed in agarose in the traditional assay, cells are captured into an agarose microwell array by gravity. The platform also expands from the size of a standard microscope slide to a 96-well format, enabling parallel processing. Here we describe the protocols of using the chip to evaluate DNA damage caused by known genotoxic agents and the cellular repair response followed after exposure. Through the integration of biological and engineering principles, this method potentiates robust and sensitive measurements of DNA damage in human cells and provides the necessary throughput for genotoxicity testing, drug development, epidemiological studies and clinical assays.

We describe here a platform that allows comet assay detection of DNA damage with unprecedented throughput. The device patterns mammalian cells into a microarray and enables parallel processing of 96 samples. The approach facilitates analysis of base level DNA damage, exposure-induced DNA damage and DNA repair kinetics.

DNA damaging agents can promote aging, disease and cancer and they are ubiquitous in the environment and produced within human cells as normal cellular ...

High Throughput Microfluidic Rapid and Low Cost Prototyping Packaging Methods

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging techniques.
The one-time use packaging technique employs UV-based and temperature curing epoxies to connect microtubes to access holes, wire-bonding for integrated circuit connections, and silver epoxy for electrical connections. This method is based on a robust assembly technique that can support relatively high pressure close to 1 psi and does not need any support to strengthen the microfluidic architecture.
Reusable packaging techniques consist of PDMS-based microtube interconnectors and anisotropic adhesive films for electrical connections. These devices are more sensitive and fragile. Consequently, Plexiglas support is added to the microfluidic structure to improve the electrical contact when anisotropic adhesive films are used, and also to strengthen the microfluidic architecture. In addition, a micromanipulator is needed to maintain tubes while using a thin PDMS layer to connect them to the access holes. Different PDMS layer thicknesses, ranging from 0.45-3 mm, are tested to compare the best adherence versus injection rates. Applied injection rates are varied from 50-300 &#956;l/hr for 0.45-3 mm PDMS layers, respectively. These techniques are mainly applicable for low-pressure applications. However, they can be extended for high-pressure ones through plasma-oxygen process to permanently seal the PDMS to glass substrates. The main advantage of this technique, besides the fact that it is reusable, consists of keeping the device observable when the microchannel length is very short (in the range of 3 mm or lower).

In this article we describe different techniques for microfluidic rapid prototyping platforms. The proposed techniques are based on ultraviolet (UV) sensitive and temperature curing epoxies, polydimethylsiloxane (PDMS) based tubing, wire-bonding, and anisotropic adhesive films. The assembling procedures presented are developed for both one-time use devices as well as reusable microfluidic systems.

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging ...

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 ...

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 ...

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly controlled, high-throughput engineered system for measuring the impact of arrayed biochemical signals on cell differentiation. Recent efforts using cell microarrays have demonstrated their utility for combinatorial studies in which many microenvironmental factors are presented in parallel. However, these efforts have focused primarily on investigating the effects of biochemical cues on cell responses. Here, we present a cell microarray platform with tunable material properties for evaluating both cell differentiation by immunofluorescence and biomechanical cell&#8211;substrate interactions by traction force microscopy. To do so, we have developed two different formats utilizing polyacrylamide hydrogels of varying Young&#39;s modulus fabricated on either microscope slides or glass-bottom Petri dishes. We provide best practices and troubleshooting for the fabrication of microarrays on these hydrogel substrates, the subsequent cell culture on microarrays, and the acquisition of data. This platform is well-suited for use in investigations of biological processes for which both biochemical (e.g., extracellular matrix composition) and biophysical (e.g., substrate stiffness) cues may play significant, intersecting roles.

Cell differentiation is regulated by a host of microenvironmental factors, including both matrix composition and substrate material properties. We describe here a technique utilizing cell microarrays in conjunction with traction force microscopy to evaluate both cell differentiation and biomechanical cell&#8211;substrate interactions as a function of microenvironmental context.

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly ...

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...

An Ultrahigh-throughput Microfluidic Platform for Single-cell Genome Sequencing

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of cellular heterogeneity in biological systems. However, single-cell sequencing of large populations has been hampered by limitations in processing genomes for sequencing. In this paper, we describe a method for single-cell genome sequencing (SiC-seq) which uses droplet microfluidics to isolate, amplify, and barcode the genomes of single cells. Cell encapsulation in microgels allows the compartmentalized purification and tagmentation of DNA, while a microfluidic merger efficiently pairs each genome with a unique single-cell oligonucleotide barcode, allowing &gt;50,000 single cells to be sequenced per run. The sequencing data is demultiplexed by barcode, generating groups of reads originating from single cells. As a high-throughput and low-bias method of single-cell sequencing, SiC-seq will enable a broader range of genomic studies targeted at diverse cell populations.

Single-cell sequencing reveals genotypic heterogeneity in biological systems, but current technologies lack the throughput necessary for the deep profiling of community composition and function. Here, we describe a microfluidic workflow for sequencing &gt;50,000 single-cell genomes from diverse cell populations.

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of ...

A Microfluidic Platform for Stimulating Chondrocytes with Dynamic Compression

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth plate cartilage architecture and results in growth modulation of long bones of children. To determine the role of compressive stress in bone growth, we created a microfluidic device actuated by pneumatic pressure, to dynamically (or statically) compress growth plate chondrocytes embedded in alginate hydrogel cylinders. In this article, we describe detailed methods for fabricating and characterizing this device. The advantages of our protocol are: 1) Five different magnitudes of compressive stress can be generated on five technical replicates in a single platform, 2) It is easy to visualize cell morphology via a conventional light microscope, 3) Cells can be rapidly isolated from the device after compression to facilitate downstream assays, and 4) The platform can be applied to study mechanobiology of any cell type that can grow in hydrogels.

This article provides detailed methods for fabricating and characterizing a pneumatically actuating microfluidic device for chondrocyte compression.

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth ...

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

Summary

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Chapters in this video

High-throughput Protein Expression Generator Using a Microfluidic Platform

High Throughput Single-cell and Multiple-cell Micro-encapsulation

CometChip: A High-throughput 96-Well Platform for Measuring DNA Damage in Microarrayed Human Cells

High Throughput Microfluidic Rapid and Low Cost Prototyping Packaging Methods

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

An Ultrahigh-throughput Microfluidic Platform for Single-cell Genome Sequencing

A Microfluidic Platform for Stimulating Chondrocytes with Dynamic Compression